The Role of MUC5B Salivary Mucin in Modulating Oral Bacterial Surface Colonization and Interspecies Competition
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The role of MUC5B salivary mucin in modulating oral bacterial surface colonization and
interspecies competition
A dissertation presented
by
Erica Shapiro Frenkel
to
The Department of Biological Sciences in Dental Medicine
in partial fulfillments of the requirements
for the degree of
Doctor of Philosophy
in the subject of
Biological Sciences in Dental Medicine
Harvard University
Cambridge, Massachusetts
March 2017
© 2017 Erica Shapiro Frenkel All rights reserved
Dissertation Advisor: Katharina Ribbeck Erica Shapiro Frenkel
The role of MUC5B salivary mucin in modulating oral bacterial surface colonization and
interspecies competition
Abstract
Mucus acts as a protective barrier on wet epithelial surfaces in the body including the gastrointestinal, respiratory, and reproductive tracts. A main component of mucus is mucin, which give mucus its viscoelastic properties and is a key component of host defense at these epithelial interfaces. Altered mucin production has been linked to gastrointestinal diseases such as Crohn’s disease and respiratory illnesses like cystic fibrosis, which highlights the importance of regulated mucin production.
Although the role of mucins in gastrointestinal and respiratory illnesses has been well characterized, little is known about how salivary mucins protect the oral cavity from common diseases such as dental caries. Streptococcus mutans is one of the primary bacteria that cause dental caries, which form when bacteria grow on tooth surfaces then produce organic acids as metabolic byproducts. The decrease in local pH leads to dissolution of tooth enamel then cavity formation. The research in this thesis uses purified human MUC5B salivary mucin to better understand how this mucin could play a role in the prevention of cavity formation. Results shown in Chapter 2 demonstrate that MUC5B significantly reduces S. mutans attachment and biofilm formation on glass and hydroxyapatite. In addition, MUC5B does not significantly reduce S. mutans viability indicating the decrease in S. mutans surface colonization is due to a shift in cells from the biofilm to the planktonic state. In Chapter 3, a dual-species competition
iii model is used to study the effects of MUC5B in a more complex environment. The two species in the model are S. mutans and Streptococcus sanguinis, which are known to compete in the oral cavity. In this study, MUC5B increases S. mutans and S. sanguinis coexistence. The reduction in interspecies competition could be caused by an increase in the relative proportion of cells in the less competitive planktonic state, which occurs in the presence of MUC5B. Taken together, the results presented in this thesis indicate that MUC5B could play a key role in protecting the oral cavity from disease and in maintaining a healthy microbiota.
iv Table of Contents
Title page……………………………………………………………………………………....…..i
Copyright page………………………………………………………………………………….....ii
Abstract…………………………………………………………………………………………...iii
Table of Contents………………………………………………………………………………….v
Acknowledgements……………………………………………………………………………….vi
List of Figures…………………………………………………………………………………...viii
Chapter I: Introduction………………………………………………………………...…………..1
Chapter II: Salivary Mucins Protect Surfaces from Colonization by Cariogenic Bacteria……...27
Chapter III: Salivary Mucins Promote the Coexistence of Competing Oral Bacterial Species….47
Chapter IV: Conclusions and Future Directions………………………………………………....63
v Acknowledgments
I would like to thank my advisor, Dr. Katharina Ribbeck, for her continual support and guidance throughout my time in her lab. She has been a wonderful mentor whose patience and encouragement allowed me to grow as a scientist and discover the world of microbiology. Words cannot express my gratitude and appreciation.
I owe a huge thank you to the Ribbeckers who have always been there for me and were truly the best labmates I could possibly imagine. Julia and Nicole K., thank you for teaching me microbiology. Both of you were especially important in my early days in the lab when I thought that chemistry was the only science I would ever need to know. Wes and Tahoura, thank you for always being there to grab coffee and making time whenever I needed to chat. Nicole B.,
Thomas, Leon, Brad, Jacob, Gerardo, Caroline, Emi, Kelsey, and Ben, -- thank you. A special thank you to Kate and Tahoura for providing feedback on my thesis drafts.
My advisory committee members, Dr. Otto Cordero, Dr. Margaret Duncan and Dr. Max
Goodson deserve a special thanks for their helpful suggestions. Also, I would like to thank my dissertation advisory committee, Dr. Margaret Duncan, Dr. Phil Stashenko, Dr. Alan
Grodzinsky, and Dr. Abigail Sloan Devlin, for their time and comments. I am grateful to the directors of the Biological Sciences in Dental Medicine program, Dr. Bjorn Olsen and Dr. Yingzi
Yang, for their encouragement.
This journey would not have been possible without the love and support I received from my family. To my Mom, Dad and brother who always helped me over the obstacles and were there to cheer for me when I succeeded, thank you. And, of course, to Eran, who always understands, knows exactly what to say and has an unwavering belief in my ability to succeed.
vi You are truly the best. Lina, thank you for never letting me forget that life is just too short to sweat the small stuff.
vii List of Figures
Figure 1.1 Mucin dysregulation in disease.
Figure 1.2 Introduction to mucins.
Figure 1.3 Potential mechanisms by which MUC5B protects the oral cavity from microbial colonization.
Figure 1.4 Mucins reduce C. albicans hyphae formation.
Figure 2.1 Sucrose enhances S. mutans attachment and biofilm formation.
Figure 2.2 Supplemental sugar alters S. mutans growth.
Figure 2.3 Salivary mucins reduce S. mutans attachment and biofilm formation.
Figure 2.4 S. mutans growth is unaffected by the presence of salivary mucins.
Figure 2.5 S. mutans survival is unaffected by salivary mucins.
Figure 2.6 Summary of conclusions.
Figure 3.1 MUC5B promotes S. mutans and S. sanguinis coexistence.
Figure 3.2 At high cell density, S. sanguinis viability decreases over time due to self-killing.
Figure 3.3 Methylcellulose does not affect S. mutans and S. sanguinis coexistence.
Figure 3.4 MUC5B reduces S. mutans and S. sanguinis surface attachment by shifting cells into the planktonic state.
viii Chapter I
Introduction
Parts of the text presented in this chapter were published in:
Frenkel ES, Ribbeck K. 2015. Salivary mucins in host defense and disease prevention. J Oral
Microbiol 7:29759.
1 Introduction
Mucins, the primary gel-forming component of mucus, provide a critical layer of protection on wet epithelial surfaces in the body including the gastrointestinal tract, female genital tract and respiratory tract. The importance of regulated mucin production in maintaining health is illustrated most clearly by studying the disease states that develop when mucin is dysregulated. Several studies have utilized knockout mice to more specifically characterize the changes that occur when mucins are not present. Mice lacking the Muc2 intestinal mucin showed signs of reduced health compared to wild type mice including increased inflammation in the distal colon, weight loss and mucosal thickening (Figure 1.1A and B) (1, 2). The authors postulate that thickening of the mucosa could be caused by a thinning of the mucus layer, which results in an increase in bacteria contacting the intestinal surface (Figure 1.1A) (1, 3). A separate study looking at respiratory function in Muc5b knockout mice showed that Muc5b deficient mice had significantly increased bacterial load in the lungs and middle ear leading to reduced survival compared to wild type mice (Figure 1.1C) (4). In the oral cavity, decreased salivary flow is linked to increased incidence of dental caries, which could be caused by reduced levels of salivary mucins (5, 6). On the other hand, mucin overproduction is associated with diseases such as asthma and cystic fibrosis, where increased mucus secretion leads to narrowing of airways (7–
9). These findings highlight the importance of regulated mucin production, but our understanding of the precise mechanisms by which mucins provide protection in the oral cavity are not well characterized.
The goal of this thesis work is to elucidate the function MUC5B plays in protecting the oral cavity using dental caries as a disease model. Streptococcus mutans, one of the primary bacteria responsible for dental caries, creates cavities when it attaches to and grows
2 Figure 1.1 wild type Muc2 -/- A.
B. C. 105 0.2
104
0.1
-actin (%) -actin 3
β 10 CFU lung per to to
0.0 102 Relative mRNA expression expression mRNA Relative wild type Muc2 -/- wild Muc5b -/- type
Figure 1.1 Mucin dysregulation in disease. (A) Colon section from wild type and Muc2 -/- mice. Bacteria are stained red and tissue is counterstained with DAPI. Double arrow represents mucus layer above epithelium in wild type mice. Arrow in Muc2 -/- mice mark bacteria. Scale bars are 100 µm. Taken from (3) with permission. Link to copyright license: https://creativecommons.org/licenses/by/4.0/legalcode. (B) TNF-alpha, an inflammatory cytokine, was significantly upregulated in the colon of Muc2 -/- mice compared to wild type mice. P = 0.03. Taken from (1) with permission. (C) Bacterial load in the lungs of wild type and Muc5b -/- mice at 6 months of age. Taken from (4) with permission.
3 on tooth surfaces. Because surface colonization is a key step in the development of caries, I first study the effect of MUC5B on S. mutans attachment, biofilm formation and survival on various surfaces. Results show that MUC5B significantly reduces S. mutans attachment and biofilm formation without bactericidal effects, indicating that MUC5B shifts cells from the biofilm into the planktonic form. I then use a dual-species competition model with S. mutans and
Streptococcus sanguinis, to determine the effect of MUC5B on interspecies interactions. In this model, MUC5B promotes S. mutans and S. sanguinis coexistence. These studies indicate that
MUC5B likely plays an important role in protecting the oral cavity from disease by modulating microbial surface colonization and interspecies competition.
The following introduction will provide an overview of what is known about the structural features of salivary mucins, potential mechanisms by which salivary mucins protect the oral cavity, and methods for studying salivary mucins.
1. Introduction to salivary mucins
There are at least 20 identified mucins throughout the human body that cover wet epithelial surfaces such as the gastrointestinal tract, respiratory tract, and eyes. A summary of areas where mucins can be found in the body is given in Figure 1.2A. Each of these mucins has a unique structure that can influence its localization and function. In addition, one type of mucin can be secreted in several different locations in the body. For example MUC5B can be found in the oral cavity, lungs and female reproductive tract (10–12). This section will address structural aspects of the mucins found in the oral cavity, MUC5B, MUC7, MUC19, MUC1, and MUC4
(13).
4 1.1 Mucins in the oral cavity
Each of the salivary mucins MUC5B, MUC7, MUC19, MUC1, and MUC4 are composed of a unique domain structure that influences the mucins’ physical properties and localization in the oral cavity (Fig. 1.2B). MUC5B is the primary gel-forming mucin in the mouth that is secreted by mucus cells in the submandibular, sublingual, palatine, and labial salivary glands (11,
14). Transcripts and glycoproteins of MUC19, another gel-forming salivary mucin, have been identified, but MUC5B is still thought to be the predominate gel-forming mucin in the oral cavity
(15–17). MUC7 is also a secreted mucin that exists primarily as monomers or dimers and lacks gel-forming properties. These monomers and dimers are able to self-associate, however, to form higher order assemblies that could facilitate bacterial aggregation (18). MUC7 localization within salivary glands varies between individuals; it has been identified in mucus cells of submandibular and sublingual glands, but the presence of MUC7 in serous cells of these glands is variable (19). MUC1 and MUC4 are membrane-associated mucins that line the ducts of parotid, submandibular, and minor salivary glands (20, 21). These mucins may play a role in cell signal transduction and could form scaffolds for secreted mucins to bind (20, 22–24). Although several salivary mucins have been introduced, the following sections will focus specifically on
MUC5B structure and function because it is the primary mucin studied in this research. MUC7 will be discussed briefly to emphasize the relevant differences between MUC5B and MUC7 protection in the oral cavity.
5 Figure 1.2
A. eyes A. nose ears oral cavity
gastrointestinal tract respiratory tract hepatobilliary system pancreas
reproductive tract
B. B. MUC5B
MUC7
MUC19
MUC1
MUC4
D domain EGF-like domain
N-glycosylation site Cystine-knot domain
Transmembrane sequence Tandem repeat
Serine/Threonine rich region cytoplasmic tail
Signal peptide SEA module
Unique sequence MUC11p15 domain
Cysteine-rich domain B domain
C domain Histatin-like sequence
Leucine-zipper Gap in sequence
6 Figure 1.2 (Continued). Introduction to mucins. (A) Epithelial surfaces where mucins can be found in the body. (B) Predicted domain structures of the salivary mucins MUC5B (42, 43), MUC7 (43, 44), MUC19 (17), MUC1 (43, 45, 46) and MUC4 (47). Note that predicted domain sequences can vary based on detection method. Figure taken from (48).
1.2 MUC5B structure and secretion
MUC5B has several unique aspects of its primary sequences that determine its ability to form gels and higher order structures. MUC5B is composed of a protein backbone with glycan chains radiating outward to form a ‘bottle-brush’ structure, which is a common characteristic of mucins. Glycan chains account for approximately 80% of MUC5B’s mass and are composed of a complex array of sugars such as fucose, galactose, and N-Acetylglucosamine (25, 26). There are several excellent papers that outline the composition of their glycan chains, which can be referred to for more detailed descriptions (25–28).
The MUC5B backbone is composed of approximately 5,700 amino acids and is broadly organized into the N-terminus, central glycosylated region, and C-terminus (29–31). The exact number of amino acids in the backbone varies among studies most likely because of variations in the tandem repeat region. There are several excellent reviews that further detail the structure of mucins (26, 32–38). In the endoplasmic reticulum, the C-terminal domain participates in disulfide bond formation, which links individual MUC5B monomers into dimers (39, 40). Then, in the golgi complex, polymer chains form through disulfide bond formation at the N-terminus and amino acids in the backbone are O-glycosylated (39, 40). MUC5B’s central glycosylated region contains repeating units of 29 amino acids that are rich in serine and threonine (41). N-
Acetylgalactosamine attaches to serine or threonine’s hydroxyl group, which provides the foundation for O-glycosylated sugar chains (26). Several excellent reviews detail the formation
7 of mucin polymers and packaging within the cell (32, 33, 49). Once the packaged mucin granule is secreted, divalent calcium ions,, which stabilize the folded mucin polymer within the secretory granule, are exchanged for monovalent sodium ions (30, 50). The increased osmotic pressure leads to hydration, which drives expansion of the polymers and formation of a gel (30). The expanded polymers cross-link via entanglement of glycoprotein polymer chains and/or non- covalent bonds formed by hydrophobic or carbohydrate-carbohydrate interactions (51–55).
Calcium may also mediate cross-linking of MUC5B to form higher order structures (56). The resulting hydrogel coats the oral epithelium as part of the protective pellicle layer and houses a vast number of oral microbes (57, 58). MUC5B’s structure and physical location in the oral cavity impact the ways in which it provides protection, which is addressed in the following section.
2. Mechanisms of protection by MUC5B
MUC5B protects the oral cavity through several different mechanisms that are influenced by its unique polymer structure. These mechanisms are summarized in Figure 1.3. First, MUC5B can interact with salivary proteins to alter their localization and retention (A), which could provide increased protection for the oral cavity. To defend the oral epithelium from potentially pathogenic microbes, MUC5B could act as a barrier (B) and/or bind microbes to facilitate their removal (C). In addition, results from this thesis demonstrate that a fourth protective mechanism is possible: MUC5B can modulate pathogenicity by facilitating bacterial dispersal (D).
8 Figure 1.3
Figure 1.3 Potential mechanisms by which MUC5B protects the oral cavity from microbial colonization. (A) MUC5B could interact with protective salivary proteins to increase their retention and broaden their localization. (B) MUC5B could form a barrier that reduces bacterial contact with the epithelial surface. (C) MUC5B could agglutinate microbes, which would facilitate their removal during swallowing. (D) MUC5B could disperse bacteria through glycan- specific interactions. Selective interactions between mucins and microbes may have downstream effects on genetic regulation that reduce virulence. Schematic not drawn to scale. Adapted from (48).
2.1 Interactions between MUC5B and salivary proteins
One way MUC5B protects the oral cavity is by binding to antibacterial salivary proteins, which can influence the proteins’ localization in the oral cavity, increase their retention time, and alter their biological activity (Figure 1.3A). Western blotting revealed that MUC5B forms heterotypic complexes with acidic and basic proline-rich proteins, statherins and histatin 1 (59).
These proteins all have antimicrobial properties; therefore, increasing their availability in saliva could be beneficial to oral health. In some cases, salivary mucins have been shown to be
9 involved in sIgA binding to the mucosal pellicle, which would enhance sIgA concentration near the oral epithelium (60). MUC5B binding to this select group of salivary proteins indicates that the formation of these complexes is protein specific (59). To better understand the nature of these complexes, Iontcheva et al. show that the interaction between MUC5B and proline-rich proteins, statherins, and histatins can be dissociated using denaturing conditions, indicating that these proteins bind through hydrophobic or ionic interactions, hydrogen bonding, or van der Waals forces (59). In some cases, proline-rich proteins and statherins were also able to form bonds with
MUC5B that were resistant to denaturing conditions, suggesting that covalent interactions may be involved in some types of complexes (59). Collectively, these studies indicate that MUC5B may serve as carriers for antibacterial salivary proteins to transport them throughout the oral cavity, increase their retention in the dental pellicle, and/or protect proteins from proteolytic degradation through the formation of complexes.
2.2 Mucins as barriers
The mucus layer that coats wet epithelia in the body acts as a barrier that protects the underlying tissue from mechanical damage, chemical insult, and pathogenic microbes (Figure
1.3B). In the oral cavity, MUC5B is the primary salivary mucin found in the mucus layer that coats soft and hard tissues, which is commonly referred to as the acquired pellicle (57, 58).
Together with saliva, the pellicle prevents desiccation and provides lubrication that protects tissues from the forces of mastication (36, 61). One of the key roles of the mucus layer is to act as a selective barrier that allows passage of certain beneficial particles, such as gases, but not potentially harmful agents such as bacteria and acids (62). In general, microbes only directly contact wet epithelium during disease states where mucus production is dysregulated, such as
10 ulcerative colitis, or when the microbes have evolved specific mechanisms to subvert the barrier.
In the stomach, for example, Helicopacter pylori increases the pH of its immediate environment by secreting urease, which reduces gastric mucus viscoelasticity and allows the pathogen to penetrate the mucus barrier and reach the gastric epithelium (63, 64). The mechanisms by which the mucus layer selects which particles and microbes can enter or pass through the barrier is an active area of research that is of great interest in the field of drug delivery. There are several excellent reviews that discuss this extensive topic (62, 65). Briefly, there are two general mechanisms by which mucins regulate the passage of particles, which include microbes. First, mucins can limit diffusion through the barrier based on the length scale of the pores formed by overlapping polymers. In general, this polymer network would allow diffusion of particles that are smaller than the pore size and exclude particles that are larger than the pore (65, 66). The biochemical properties of mucins are another way that they can impart selective diffusion characteristics to the mucus barrier (67–69). Sialic acid and sulfate residues on mucin give the polymer a net negative charge, which can exclude particles based on their chemical properties
(70, 71). Mucin polymers can also participate in hydrogen bonding and hydrophilic interactions that can limit or increase the ability of particles to penetrate the mucus layer depending on the specific properties of the particles (70, 71).
2.3 Mucins binding microbes
In addition to the structural characteristics that allow mucins to form a selective barrier, mucins also contain properties that allow them to directly interact with microbes (Figure 1.3C).
Early studies show that submandibular/sublingual gland saliva and salivary mucins aggregate specific strains of suspended bacteria and induce bacterial attachment to mucin-coated surfaces
11 (72–75). These studies primarily focus on interactions between salivary mucins and oral streptococci. Because mucins induce aggregation or surface attachment of certain bacterial species, this indicates that the bacteria recognize and bind specific glycans on the mucins, such as sialic acid and blood-group antigens (75–77). The increase in surface attachment could be beneficial because sugars in mucins’ glycan chains could act as decoys for the epithelial surface
(13). Importantly, many of these early studies do not distinguish between MUC5B and MUC7 salivary mucins. When these two mucins are electrophoretically separated, however, radiolabeled Streptococcus sanguinis, S. sobrinus, and S. oralis bind to MUC7 but not MUC5B
(78). Another similar study also shows that MUC7 aggregates S. gordonii and promotes its adherence to surfaces whereas MUC5B has no effect on aggregation or binding (79). These findings indicate that MUC7 and MUC5B exert their protective effects through different mechanisms.
2.4 MUC5B mediated reduction in microbial pathogenicity
In contrast to MUC7, MUC5B appears to bind only a limited number of oral bacteria despite its heterogeneous glycan chains. Murray et al. tested the binding of 16 species of streptococci to MUC5B, but none of the tested strains bound this mucin (78). One explanation is that the heterogeneous glycan chains found on MUC5B would prevent binding due to the inability of bacteria to form multiple bonds or attachment points. In line with this hypothesis,
Haemophilus parainfluenzae was shown to bind MUC5B, but this bacterium interacts with the naked peptide backbone as opposed to the glycan chains (80). Helicobacter pylori is another bacterium that binds MUC5B through a neutrophil-activating protein on its surface that mediates binding to sulfated glycans (81). Although few studies have shown bacteria binding to MUC5B,
12 there could be other oral microbes that do interact directly with MUC5B, but their interactions have not yet been characterized.
The limited number of bacteria known to bind MUC5B compared to MUC7 highlights the point that MUC5B protects the oral cavity in a unique way. The research presented in this thesis, in addition to other studies from the Ribbeck lab, indicate that mucins promote bacterial dispersal, which may have downstream effects that lead to a reduction in microbial virulence
(Figure 1.3D). A recent study from the Ribbeck lab shows that Muc5ac gastric mucin reduces
Candida albicans surface colonization by shifting cells away from the aggregated biofilm into the planktonic state (82). In addition, results show that MUC5B and Muc5ac repress C. albicans virulence by reducing the formation of hyphae, which are associated with host cell invasion
(Figure 1.4) (82–85). C. albicans is the primary microbe responsible for oral candidiasis, an overgrowth of the fungus on oral tissues. The opportunistic fungus exists as part of the normal oral flora in many individuals but can become pathogenic in immune compromised individuals and lead to life-threatening systemic infection if left unchecked (86, 87). MUC5B’s ability to reduce C. albicans virulence without killing the fungus could explain how this opportunistic pathogen can exist as part of a healthy oral microbiota without the development of overt candidiasis. MUC5B’s ability to form a gel layer that guards against pathogenic microbes but does not cause bacterial killing is a unique property that contrasts with other defense proteins in saliva, such as antimicrobial peptides.
13 Figure 1.4
− Mucin Muc5ac MUC5B
Figure 1.4 Mucins reduce C. albicans hyphae formation. Phase-contrast image of C. albicans after growth in the absence of mucins, in Muc5ac pig gastric mucin and in MUC5B human salivary mucin. Taken from (82) with permission. Link to license: https://creativecommons.org/licenses/by-nc-sa/3.0/legalcode.
3. Methods for studying salivary mucins
Because salivary mucins are relatively accessible compared to other types of mucins, such as gastric or intestinal mucin, studies using salivary mucins generally rely on purified human mucins or commercially available bovine submaxillary mucin. In contrast, studies on less accessible types of mucins commonly use animal organs for mucin purification, commercial mucin sources or mucus-secreting cell lines. Although commercially available submaxillary mucin is relatively easy to obtain compared to purification of salivary mucin from whole saliva, the properties of commercial mucin do not mimic those of purified native mucin. Specifically, commercial mucin does not maintain its gel-forming properties, which is one of the central characteristics of the mucin family of glycoproteins (63, 88). In addition, studies have shown that commercially purified mucins can be cytotoxic and do not possess the same protective properties as natively purified mucins (89). For example, commercial Muc5ac porcine gastric mucin is less efficient at reducing HPV-16 infection of HeLa cells compared to natively purified Muc5ac (89).
14 During the commercial purification process mucins may be proteolytically degraded, which reduces their ability to form gels and alters their chemical properties (90).
Because of the limitations of commercially available salivary mucins, I developed a protocol to purify MUC5B from human saliva. Briefly, the protocol involves collecting saliva that is enriched for sublingual and submandibular gland secretions (the primary sources of
MUC5B), centrifugation to remove debris, then size exclusion chromatography. This purification protocol is relatively gentle and leaves mucins in their native form (91, 92). One drawback of this type of set up, however, is that it does not account for the other components of saliva and the mucus barrier that may function with MUC5B to protect the oral cavity. In my research, I study these purified mucins in a three-dimensional environment as opposed to a surface coating. The cell-repellent properties of mucin coatings have been studied, but these coatings may not accurately represent the environment in the oral cavity (93–95). By studying MUC5B in a three- dimensional environment, mucin polymers are fully expanded and domains can interact to form an extended gel network, which, I hypothesize, is more similar to the environment in the oral cavity (30, 34, 96, 97). By using purified human MUC5B, I am able to evaluate the effects of this mucin in a defined environment that is easily manipulated and where experimental results can be readily interpreted.
To better understand how MUC5B affects microbes to protect the oral cavity, I study
MUC5B in the context of a specific disease model – dental caries. Caries is the most common chronic childhood disease and approximately 100% of adults have dental caries, therefore this model is highly relevant (98). In Chapter 2, I use purified human MUC5B mucin to study how it
15 affects Streptococcus mutans UA159 attachment and biofilm formation. Although there are several bacteria that have recently been implicated in the formation of dental caries, S. mutans is considered to be one of the primary cavity-causing bacteria. Results indicate that MUC5B significantly reduces S. mutans attachment and biofilm formation, which are key steps in the process of cavity formation. Furthermore, I show that MUC5B does not alter S. mutans growth rate or have bactericidal effects indicating that MUC5B reduces surface colonization by shifting cells from the biofilm into the planktonic state.
In Chapter 3, I designed an experimental dual-species model to study the effects of
MUC5B on bacteria in a more complex environment. I use the model to determine the influence of MUC5B on interspecies competition using bacterial viability as a readout. The model is composed of S. mutans UA159 and S. sanguinis JFP36, which are known to compete in the oral cavity through the secretion of antimicrobial peptides and hydrogen peroxide, respectively (99–
102). Results indicate that MUC5B increases coexistence of the two bacteria and shifts cells from the mixed-species biofilm into the less competitive planktonic form (103, 104). Based on these findings, I hypothesize that MUC5B modulates oral bacteria in ways that reduce bacterial surface colonization and promote coexistence in the oral microbiota.
When the findings of this thesis are viewed as a whole, a trend begins to emerge that hints at a general underlying mechanism of protection: MUC5B protects the oral cavity by suppressing virulent traits, such as bacterial surface attachment, without killing the organism.
This mechanism of protection is very different than the classical view of mucus as a simple physical barrier overlying the epithelium. In addition, this work emphasizes that the mechanism through which MUC5B protects the oral cavity is distinct from MUC7 salivary mucins, which primarily directly bind microbes (78, 105, 106). MUC5B’s ability to allow opportunistic
16 pathogens to exist and live within the oral microbiota as non-pathogenic residents is unprecedented by other antimicrobial proteins in saliva. This work illustrating that mucins influence bacterial surface colonization and interspecies interactions enhances our understanding of oral disease pathogenesis and could lead to novel strategies for disease prevention and treatment.
17 References
1. Van der Sluis M, De Koning B, De Bruijn A, Velcich A, Meijerink J, Van Goudoever J, Büller H, Dekker J, Van Seuningen I, Renes I, Einerhand A. 2006. Muc2-deficient mice spontaneously develop colitis, indicating that MUC2 is critical for colonic protection. Gastroenterology 131:117–29.
2. Bergstrom KSB, Kissoon-Singh V, Gibson DL, Ma C, Montero M, Sham HP, Ryz N, Huang T, Velcich A, Finlay BB, Chadee K, Vallance BA. 2010. Muc2 Protects against Lethal Infectious Colitis by Disassociating Pathogenic and Commensal Bacteria from the Colonic Mucosa. PLoS Pathog 6:e1000902.
3. Wenzel UA, Magnusson MK, Rydström A, Jonstrand C, Hengst J, Johansson ME, Velcich A, Öhman L, Strid H, Sjövall H, Hansson GC, Wick MJ. 2014. Spontaneous Colitis in Muc2-Deficient Mice Reflects Clinical and Cellular Features of Active Ulcerative Colitis. PLOS ONE 9:e100217.
4. Roy MG, Livraghi-Butrico A, Fletcher AA, McElwee MM, Evans SE, Boerner RM, Alexander SN, Bellinghausen LK, Song AS, Petrova YM, Tuvim MJ, Adachi R, Romo I, Bordt AS, Bowden MG, Sisson JH, Woodruff PG, Thornton DJ, Rousseau K, De la Garza MM, Moghaddam SJ, Karmouty-Quintana H, Blackburn MR, Drouin SM, Davis CW, Terrell KA, Grubb BR, O’Neal WK, Flores SC, Cota-Gomez A, Lozupone CA, Donnelly JM, Watson AM, Hennessy CE, Keith RC, Yang IV, Barthel L, Henson PM, Janssen WJ, Schwartz DA, Boucher RC, Dickey BF, Evans CM. 2014. Muc5b is required for airway defence. Nature 505:412–416.
5. Brown LR, Dreizen S, Handler S, Johnston DA. 1975. Effect of Radiation-Induced Xerostomia on Human Oral Microflora. J Dent Res 54:740–750.
6. Dreizen S, Brown LR, Daly TE, Drane JB. 1977. Prevention of Xerostomia-Related Dental Caries in Irradiated Cancer Patients. J Dent Res 56:99–104.
7. Rose MC, Voynow JA. 2006. Respiratory tract mucin genes and mucin glycoproteins in health and disease. Physiol Rev 86:245–278.
8. Kreda SM, Davis CW, Rose MC. 2012. CFTR, Mucins, and Mucus Obstruction in Cystic Fibrosis. Cold Spring Harb Perspect Med 2:a009589.
9. Green TD, Crews AL, Park J, Fang S, Adler KB. 2011. Regulation of mucin secretion and inflammation in asthma; A role for MARCKS protein? Biochim Biophys Acta 1810:1110–1113.
10. Andersch-Björkman Y, Thomsson KA, Larsson JMH, Ekerhovd E, Hansson GC. 2007. Large Scale Identification of Proteins, Mucins, and Their O-Glycosylation in the Endocervical Mucus during the Menstrual Cycle. Mol Cell Proteomics 6:708–716.
18 11. Thornton DJ, Khan N, Mehrotra R, Howard M, Sheehan JK, Veerman E, Packer NH. 1999. Salivary mucin MG1 is comprised almost entirely of different glycosylated forms of the MUC5B gene product. Glycobiology 9:293–302.
12. Wickström C, Davies JR, Eriksen GV, Veerman EC, Carlstedt I. 1998. MUC5B is a major gel-forming, oligomeric mucin from human salivary gland, respiratory tract and endocervix: identification of glycoforms and C-terminal cleavage. Biochem J 334:685– 693.
13. Linden SK, Sutton P, Karlsson NG, Korolik V, McGuckin MA. 2008. Mucins in the mucosal barrier to infection. Mucosal Immunol 1:183–197.
14. Nielsen PA, Bennett EP, Wandall HH, Therkildsen MH, Hannibal J, Clausen H. 1997. Identification of a major human high molecular weight salivary mucin (MG1) as tracheobronchial mucin MUC5B. Glycobiology 7:413–419.
15. Culp DJ, Robinson B, Cash MN, Bhattacharyya I, Stewart C, Cuadra-Saenz G. 2015. Salivary Mucin 19 Glycoproteins: Innate Immune Functions in Streptococcus mutans- Induced Caries in Mice and Evidence for Expression in Human Saliva. J Biol Chem 290:2993-3008.
16. Rousseau K, Kirkham S, Johnson L, Fitzpatrick B, Howard M, Adams EJ, Rogers DF, Knight D, Clegg P, Thornton DJ. 2008. Proteomic analysis of polymeric salivary mucins: no evidence for MUC19 in human saliva. Biochem J 413:545–552.
17. Chen Y, Zhao YH, Kalaslavadi TB, Hamati E, Nehrke K, Le AD, Ann DK, Wu R. 2004. Genome-wide search and identification of a novel gel-forming mucin MUC19/Muc19 in glandular tissues. Am J Respir Cell Mol Biol 30:155–165.
18. Mehrotra R, Thornton DJ, Sheehan JK. 1998. Isolation and physical characterization of the MUC7 (MG2) mucin from saliva: evidence for self-association. Biochem J 334:415– 422.
19. Piludu M, Rayment SA, Liu B, Offner GD, Oppenheim FG, Troxler RF, Hand AR. 2003. Electron Microscopic Immunogold Localization of Salivary Mucins MG1 and MG2 in Human Submandibular and Sublingual Glands. J Histochem Cytochem 51:69–79.
20. Liu B, Lague JR, Nunes DP, Toselli P, Oppenheim FG, Soares RV, Troxler RF, Offner GD. 2002. Expression of Membrane-associated Mucins MUC1 and MUC4 in Major Human Salivary Glands. J Histochem Cytochem 50:811–820.
21. Sengupta A, Valdramidou D, Huntley S, Hicks SJ, Carrington SD, Corfield AP. 2001. Distribution of MUC1 in the normal human oral cavity is localized to the ducts of minor salivary glands. Arch Oral Biol 46:529–538.
19 22. Offner GD, Troxler RF. 2000. Heterogeneity of High-molecular-weight Human Salivary Mucins. Adv Dent Res 14:69–75.
23. Gabryel-Porowska H, Gornowicz A, Bielawska A, Wójcicka A, Maciorkowska E, Grabowska SZ, Bielawski K. 2014. Mucin levels in saliva of adolescents with dental caries. Med Sci Monit 20:72–77.
24. Parmley RR, Gendler SJ. 1998. Cystic fibrosis mice lacking Muc1 have reduced amounts of intestinal mucus. J Clin Invest 102:1798–1806.
25. Thomsson KA, Prakobphol A, Leffler H, Reddy MS, Levine MJ, Fisher SJ, Hansson GC. 2002. The salivary mucin MG1 (MUC5B) carries a repertoire of unique oligosaccharides that is large and diverse. Glycobiology 12:1–14.
26. Strous GJ, Dekker J. 1992. Mucin-Type Glycoproteins. Crit Rev Biochem Mol Biol 27:57–92.
27. Amerongen AVN, Bolscher JGM, Veerman ECI. 1995. Salivary mucins: protective functions in relation to their diversity. Glycobiology 5:733–740.
28. Zalewska A, Zwierz K, Zółkowski K, Gindzieński A. 2000. Structure and biosynthesis of human salivary mucins. Acta Biochim Pol 47:1067–1079.
29. Desseyn JL, Buisine MP, Porchet N, Aubert JP, Laine A. 1998. Genomic organization of the human mucin gene MUC5B. cDNA and genomic sequences upstream of the large central exon. J Biol Chem 273:30157–30164.
30. Kesimer M, Makhov AM, Griffith JD, Verdugo P, Sheehan JK. 2010. Unpacking a gel- forming mucin: a view of MUC5B organization after granular release. Am J Physiol - Lung Cell Mol Physiol 298:L15–L22.
31. Preciado D, Goyal S, Rahimi M, Watson AM, Brown KJ, Hathout Y, Rose MC. 2010. MUC5B is the predominant mucin glycoprotein in Chronic Otitis Media Fluid. Pediatr Res 68:231–236.
32. Perez-Vilar J, Hill RL. 1999. The Structure and Assembly of Secreted Mucins. J Biol Chem 274:31751–31754.
33. Perez-Vilar J, Mabolo R. 2007. Gel-forming mucins. Notions from in vitro studies. Histol Histopathol 22:455–464.
34. Bansil R, Turner BS. 2006. Mucin structure, aggregation, physiological functions and biomedical applications. Curr Opin Colloid Interface Sci 11:164–170.
35. Tabak LA. 1990. Structure And Function of Human Salivary Mucins. Crit Rev Oral Biol Med 1:229–234.
20 36. Tabak LA, Levine MJ, Mandel ID, Ellison SA. 1982. Role of salivary mucins in the protection of the oral cavity. J Oral Pathol 11:1–17.
37. Tabak LA. 1995. In Defense of the Oral Cavity: Structure, Biosynthesis, and Function of Salivary Mucins. Annu Rev Physiol 57:547–564.
38. Levine MJ, Reddy MS, Tabak LA, Loomis RE, Bergey EJ, Jones PC, Cohen RE, Stinson MW, Al-Hashimi I. 1987. Structural Aspects of Salivary Glycoproteins. J Dent Res 66:436–441.
39. Perez-Vilar J, Eckhardt AE, DeLuca A, Hill RL. 1998. Porcine Submaxillary Mucin Forms Disulfide-linked Multimers through Its Amino-terminal D-domains. J Biol Chem 273:14442–14449.
40. Perez-Vilar J, Eckhardt AE, Hill RL. 1996. Porcine Submaxillary Mucin Forms Disulfide- bonded Dimers between Its Carboxyl-terminal Domains. J Biol Chem 271:9845–9850.
41. Desseyn JL, Aubert JP, Van Seuningen I, Porchet N, Laine A. 1997. Genomic organization of the 3’ region of the human mucin gene MUC5B. J Biol Chem 272:16873– 16883.
42. Desseyn J-L, Aubert J-P, Porchet N, Laine A. 2000. Evolution of the Large Secreted Gel- Forming Mucins. Mol Biol Evol 17:1175–1184.
43. Andrianifahanana M, Moniaux N, Batra SK. 2006. Regulation of mucin expression: Mechanistic aspects and implications for cancer and inflammatory diseases. Biochim Biophys Acta 1765:189–222.
44. Gururaja TL, Ramasubbu N, Venugopalan P, Reddy MS, Ramalingam K, Levine MJ. 1998. Structural features of the human salivary mucin, MUC7. Glycoconj J 15:457–467.
45. Moniaux N, Escande F, Porchet N, Aubert JP, Batra SK. 2001. Structural organization and classification of the human mucin genes. Front Biosci 6:D1192–1206.
46. Bork P, Patthy L. 1995. The SEA module: a new extracellular domain associated with O- glycosylation. Protein Sci 4:1421–1425.
47. Choudhury A, Moniaux N, Ringel J, King J, Moore E, Aubert J-P, Batra SK. 2001. Alternate splicing at the 3′-end of the human pancreatic tumor-associated mucin MUC4 cDNA. Teratog Carcinog Mutagen 21:83–96.
48. Frenkel ES, Ribbeck K. 2015. Salivary mucins in host defense and disease prevention. J Oral Microbiol 7:29759.
49. Adler KB, Tuvim MJ, Dickey BF. 2013. Regulated mucin secretion from airway epithelial cells. Neuroendocr Sci 4:129.
21 50. Ridley C, Kouvatsos N, Raynal BD, Howard M, Collins RF, Desseyn J-L, Jowitt TA, Baldock C, Davis CW, Hardingham TE, Thornton DJ. 2014. Assembly of the Respiratory Mucin MUC5B: a new model for a gel-forming mucin. J Biol Chem 289:16409–16420.
51. Perez-Vilar J. 2007. Mucin Granule Intraluminal Organization. Am J Respir Cell Mol Biol 36:183–190.
52. Verdugo P. 1990. Goblet Cells Secretion and Mucogenesis. Annu Rev Physiol 52:157– 176.
53. Bromberg LE, Barr DP. 2000. Self-Association of Mucin. Biomacromolecules 1:325–334.
54. Soby LM, Jamieson AM, Blackwell J, Jentoft N. 1990. Viscoelastic properties of solutions of ovine submaxillary mucin. Biopolymers 29:1359–1366.
55. McCullagh CM, Jamieson AM, Blackwell J, Gupta R. 1995. Viscoelastic properties of human tracheobronchial mucin in aqueous solution. Biopolymers 35:149–159.
56. Raynal BDE, Hardingham TE, Sheehan JK, Thornton DJ. 2003. Calcium-dependent protein interactions in MUC5B provide reversible cross-links in salivary mucus. J Biol Chem 278:28703–28710.
57. Al-Hashimi I, Levine MJ. 1989. Characterization of in vivo salivary-derived enamel pellicle. Arch Oral Biol 34:289–295.
58. Cárdenas M, Elofsson U, Lindh L. 2007. Salivary Mucin MUC5B Could Be an Important Component of in Vitro Pellicles of Human Saliva: An in Situ Ellipsometry and Atomic Force Microscopy Study. Biomacromolecules 8:1149–1156.
59. Iontcheva I, Oppenheim FG, Troxler RF. 1997. Human salivary mucin MG1 selectively forms heterotypic complexes with amylase, proline-rich proteins, statherin, and histatins. J Dent Res 76:734–743.
60. Gibbins HL, Proctor GB, Yakubov GE, Wilson S, Carpenter GH. 2015. SIgA Binding to Mucosal Surfaces Is Mediated by Mucin-Mucin Interactions. PLoS ONE 10:e0119677.
61. Hughes CV, Baum BJ, Fox PC, Marmary Y, Yeh C-K, Sonies BC. 1987. Oral-pharyngeal dysphagia: A common sequela of salivary gland dysfunction. Dysphagia 1:173–177.
62. Cone RA. 2009. Barrier properties of mucus. Adv Drug Deliv Rev 61:75–85.
63. Celli JP, Turner BS, Afdhal NH, Ewoldt RH, McKinley GH, Bansil R, Erramilli S. 2007. Rheology of Gastric Mucin Exhibits a pH-Dependent Sol−Gel Transition. Biomacromolecules 8:1580–1586.
22 64. Celli JP, Turner BS, Afdhal NH, Keates S, Ghiran I, Kelly CP, Ewoldt RH, McKinley GH, So P, Erramilli S, Bansil R. 2009. Helicobacter pylori moves through mucus by reducing mucin viscoelasticity. Proc Natl Acad Sci 106:14321–14326.
65. Lai SK, Wang Y-Y, Hanes J. 2009. Mucus-penetrating nanoparticles for drug and gene delivery to mucosal tissues. Adv Drug Deliv Rev 61:158–171.
66. Lieleg O, Vladescu I, Ribbeck K. 2010. Characterization of particle translocation through mucin hydrogels. Biophys J 98:1782–1789.
67. Olmsted SS, Padgett JL, Yudin AI, Whaley KJ, Moench TR, Cone RA. 2001. Diffusion of macromolecules and virus-like particles in human cervical mucus. Biophys J 81:1930– 1937.
68. Li L, Lieleg O, Jang S, Ribbeck K, Han J. 2012. A microfluidic in vitro system for the quantitative study of the stomach mucus barrier function. Lab Chip 12:4071–4079.
69. Li LD, Crouzier T, Sarkar A, Dunphy L, Han J, Ribbeck K. 2013. Spatial Configuration and Composition of Charge Modulates Transport into a Mucin Hydrogel Barrier. Biophys J 105:1357–1365.
70. Rogunova MA, Blackwell J, Jamieson AM, Pasumar-Thy M, Gerken TA. 1997. Effects of lipid on the structure and rheology of gels formed by canine submaxillary mucin. Biorheology 34:295–308.
71. Harding SE, Davis SSB, Deacon MP, Fiebrig I. 1999. Biopolymer Mucoadhesives. Biotechnol Genet Eng Rev 16:41–86.
72. Koop HM, Valentijn-Benz M, Nieuw Amerongen AV, Roukema PA, de Graaff J. 1990. Involvement of human mucous saliva and salivary mucins in the aggregation of the oral bacteria Streptococcus sanguis, Streptococcus oralis, and Streptococcus rattus. Antonie Van Leeuwenhoek 57:245–252.
73. Gibbons RJ, Cohen L, Hay DI. 1986. Strains of Streptococcus mutans and Streptococcus sobrinus attach to different pellicle receptors. Infect Immun 52:555–561.
74. Gibbons RJ, Qureshi JV. 1978. Selective binding of blood group-reactive salivary mucins by Streptococcus mutans and other oral organisms. Infect Immun 22:665–671.
75. Levine MJ, Herzberg MC, Levine MS, Ellison SA, Stinson MW, Li HC, van Dyke T. 1978. Specificity of salivary-bacterial interactions: role of terminal sialic acid residues in the interaction of salivary glycoproteins with Streptococcus sanguis and Streptococcus mutans. Infect Immun 19:107–115.
23 76. Williams RC, Gibbons RJ. 1975. Inhibition of streptococcal attachment to receptors on human buccal epithelial cells by antigenically similar salivary glycoproteins. Infect Immun 11:711–718.
77. Murray PA, Levine MJ, Tabak LA, Reddy MS. 1982. Specificity of salivary-bacterial interactions: II. Evidence for a lectin on Streptococcus sanguis with specificity for a NeuAc alpha 2, 3Ga1 beta 1, 3Ga1NAc sequence. Biochem Biophys Res Commun 106:390–396.
78. Murray PA, Prakobphol A, Lee T, Hoover CI, Fisher SJ. 1992. Adherence of oral streptococci to salivary glycoproteins. Infect Immun 60:31–38.
79. Ligtenberg AJ, Walgreen-Weterings E, Veerman EC, de Soet JJ, de Graaff J, Amerongen AV. 1992. Influence of saliva on aggregation and adherence of Streptococcus gordonii HG 222. Infect Immun 60:3878–3884.
80. Veerman EC, Ligtenberg AJ, Schenkels LC, Walgreen-Weterings E, Nieuw Amerongen AV. 1995. Binding of human high-molecular-weight salivary mucins (MG1) to Hemophilus parainfluenzae. J Dent Res 74:351–357.
81. Namavar F, Sparrius M, Veerman EC, Appelmelk BJ, Vandenbroucke-Grauls CM. 1998. Neutrophil-activating protein mediates adhesion of Helicobacter pylori to sulfated carbohydrates on high-molecular-weight salivary mucin. Infect Immun 66:444–447.
82. Kavanaugh NL, Zhang AQ, Nobile CJ, Johnson AD, Ribbeck K. 2014. Mucins Suppress Virulence Traits of Candida albicans. mBio 5:e01911–14.
83. Ogasawara A, Komaki N, Akai H, Hori K, Watanabe H, Watanabe T, Mikami T, Matsumoto T. 2007. Hyphal formation of Candida albicans is inhibited by salivary mucin. Biol Pharm Bull 30:284–286.
84. Dalle F, Wächtler B, L’Ollivier C, Holland G, Bannert N, Wilson D, Labruère C, Bonnin A, Hube B. 2010. Cellular interactions of Candida albicans with human oral epithelial cells and enterocytes. Cell Microbiol 12:248–271.
85. Phan QT, Belanger PH, Filler SG. 2000. Role of Hyphal Formation in Interactions of Candida albicans with Endothelial Cells. Infect Immun 68:3485–3490.
86. Klein RS, Harris CA, Small CB, Moll B, Lesser M, Friedland GH. 1984. Oral Candidiasis in High-Risk Patients as the Initial Manifestation of the Acquired Immunodeficiency Syndrome. N Engl J Med 311:354–358.
87. Schelenz S. 2008. Management of candidiasis in the intensive care unit. J Antimicrob Chemother 61:i31–i34.
24 88. Kočevar-Nared J, Kristl J, Šmid-Korbar J. 1997. Comparative rheological investigation of crude gastric mucin and natural gastric mucus. Biomaterials 18:677–681.
89. Lieleg O, Lieleg C, Bloom J, Buck CB, Ribbeck K. 2012. Mucin biopolymers as broad- spectrum antiviral agents. Biomacromolecules 13:1724–1732.
90. Bhaskar KR, Gong DH, Bansil R, Pajevic S, Hamilton JA, Turner BS, LaMont JT. 1991. Profound increase in viscosity and aggregation of pig gastric mucin at low pH. Am J Physiol 261:G827–G832.
91. J. Schömig V, T. Käsdorf B, Scholz C, Bidmon K, Lieleg O, Berensmeier S. 2016. An optimized purification process for porcine gastric mucin with preservation of its native functional properties. RSC Adv 6:44932–44943.
92. Davies JR, Wickström C, Thornton DJ. 2012. Gel-forming and cell-associated mucins: preparation for structural and functional studies. Methods Mol Biol 842:27–47.
93. Bushnak IA, Labeed FH, Sear RP, Keddie JL. 2010. Adhesion of microorganisms to bovine submaxillary mucin coatings: effect of coating deposition conditions. Biofouling 26:387–397.
94. Shi L, Ardehali R, Caldwell KD, Valint P. 2000. Mucin coating on polymeric material surfaces to suppress bacterial adhesion. Colloids Surf B Biointerfaces 17:229–239.
95. Crouzier T, Jang H, Ahn J, Stocker R, Ribbeck K. 2013. Cell Patterning with Mucin Biopolymers. Biomacromolecules 14:3010–3016.
96. Cao X, Bansil R, Bhaskar KR, Turner BS, LaMont JT, Niu N, Afdhal NH. 1999. pH- dependent conformational change of gastric mucin leads to sol-gel transition. Biophys J 76:1250–1258.
97. Yakubov GE, Papagiannopoulos A, Rat E, Easton RL, Waigh TA. 2007. Molecular Structure and Rheological Properties of Short-Side-Chain Heavily Glycosylated Porcine Stomach Mucin. Biomacromolecules 8:3467–3477.
98. World Health Organization. Fact Sheet N318 - Oral Health. 2012.
99. Kreth J, Merritt J, Shi W, Qi F. 2005. Competition and Coexistence between Streptococcus mutans and Streptococcus sanguinis in the Dental Biofilm. J Bacteriol 187:7193–7203.
100. Kreth J, Zhang Y, Herzberg MC. 2008. Streptococcal Antagonism in Oral Biofilms: Streptococcus sanguinis and Streptococcus gordonii Interference with Streptococcus mutans. J Bacteriol 190:4632–4640.
25 101. Ge X, Kitten T, Chen Z, Lee SP, Munro CL, Xu P. 2008. Identification of Streptococcus sanguinis Genes Required for Biofilm Formation and Examination of Their Role in Endocarditis Virulence. Infect Immun 76:2551–2559.
102. Senty Turner L, Das S, Kanamoto T, Munro CL, Kitten T. 2009. Development of genetic tools for in vivo virulence analysis of Streptococcus sanguinis. Microbiology 155:2573– 2582.
103. Oliveira NM, Martinez-Garcia E, Xavier J, Durham WM, Kolter R, Kim W, Foster KR. 2015. Biofilm Formation As a Response to Ecological Competition. PLoS Biol 13:e1002191.
104. Schluter J, Nadell CD, Bassler BL, Foster KR. 2015. Adhesion as a weapon in microbial competition. ISME J 9:139–149.
105. Moshier A, Reddy MS, Scannapieco FA. 1996. Role of type 1 fimbriae in the adhesion of Escherichia coli to salivary mucin and secretory immunoglobulin A. Curr Microbiol 33:200–208.
106. Heo S-M, Choi K-S, Kazim LA, Reddy MS, Haase EM, Scannapieco FA, Ruhl S. 2013. Host defense proteins derived from human saliva bind to Staphylococcus aureus. Infect Immun 81:1364–1373.
26 Chapter II
Salivary Mucins Protect Surfaces from Colonization by Cariogenic Bacteria
Results presented in this chapter were published in:
Frenkel ES, Ribbeck K. 2015. Salivary Mucins Protect Surfaces from Colonization by
Cariogenic Bacteria. Appl Environ Microbiol 81:332–338.
27 Abstract
Understanding how the body’s natural defenses function to protect the oral cavity from the myriad of bacteria that colonize its surfaces is an ongoing topic of research that can lead to breakthroughs in treatment and prevention. This study focuses on elucidating the connection between MUC5B salivary mucins and dental caries, one of the most common oral diseases.
Dental caries is predominantly caused by Streptococcus mutans attachment and biofilm formation on the tooth surface. Once S. mutans attaches to the tooth, it produces organic acids as metabolic by-products that dissolve tooth enamel, leading to cavity formation. We utilize CFU counts and fluorescence microscopy to quantitatively show that S. mutans attachment and biofilm formation are most robust in the presence of sucrose and that aqueous solutions of purified human MUC5B protect surfaces by acting as an antibiofouling agent in the presence of sucrose. In addition, we find that MUC5B does not alter S. mutans growth and decreases surface attachment and biofilm formation by maintaining S. mutans in the planktonic form. These insights point to the importance of salivary mucins in oral health and lead to a better understanding of how MUC5B could play a role in cavity prevention or diagnosis.
Introduction
The viscoelastic properties of mucus are attributed to mucins, large glycoproteins that play a key role in host defense and maintaining a healthy microbial environment (1–3). The diseases that result from dysregulated mucin production highlight the necessity of mucins as one of the body’s key natural defenses; however, few studies have focused specifically on the connection between MUC5B salivary mucins and oral diseases. This study fills this gap in understanding by exploring the connection between purified human MUC5B and the virulence of
28 Streptococcus mutans (4). MUC7 is another salivary mucin, but MUC5B is the primary mucin component of the dental pellicle coating the soft and hard tissues in the oral cavity (5, 6).
S. mutans is a biofilm-forming facultative anaerobic bacterium that produces three glucosyltransferase enzymes to synthesize glucans from dietary sugar (7–9). Glucans are sticky polymers that allow the cells to attach to the tooth surface and form an extracellular matrix that protects it from host defenses and mechanical removal (10, 11). Once S. mutans attaches to the tooth surface, organic acids, which are produced as metabolic by-products, become concentrated within the extracellular matrix and cause a drop in pH from neutral to 5 or below. This acidic environment begins dissolving tooth enamel, leading to cavity formation, and the high tolerance of S. mutans for acidic environments gives it an ecological advantage. Without proper hygiene and nutritional awareness, S. mutans can proliferate quickly, causing serious damage to the tooth structure. S. mutans biofilm formation is particularly problematic in the interproximal spaces between teeth, where mechanical removal is difficult.
Because S. mutans attachment and biofilm formation are critical steps in cavity formation, we use CFU counts and fluorescence microscopy to quantify the effects of supplemental sugar and purified human salivary MUC5B on these key stages of disease progression. We first validate our mucin studies by showing that S. mutans attachment and biofilm formation are most robust in the presence of sucrose as opposed to glucose. When supplemental MUC5B is added in the presence of sucrose, however, S. mutans attachment and biofilm formation are significantly decreased. Although the number of surface-attached bacteria decreases in the presence of
MUC5B, we show that bacterial growth is unchanged in the presence of MUC5B and the observed effects are due to increased numbers of S. mutans cells in the planktonic form. These findings that link MUC5B with a reduction in S. mutans surface colonization could significantly
29 impact our understanding of the pathogenesis of cavity formation and aid in the development of novel oral diagnostic methods or strategies for disease prevention.
Results and Discussion
Sucrose enhances S. mutans attachment and biofilm formation.
To determine the growth conditions where S. mutans attachment and biofilm formation are most robust, we investigated the effect of the addition of sucrose or glucose to BHI medium.
The role of sucrose in enhancing S. mutans biofilm formation has been well established using genetic analysis and biochemical assays studying biofilm architecture (12–15). Here we characterized this phenomenon using a quantitative method that directly evaluates the number of live S. mutans cells attached and producing biofilm on various surfaces. S. mutans was inoculated into BHI containing 1% sucrose or 1% glucose. CFU counts were used to evaluate attachment at 20, 40, and 60 min and biofilm formation at 6, 18, and 24 h. Attachment was defined to occur at time points up to 60 min because the doubling time of S. mutans in exponential phase is approximately 1.5 h. Experiments were carried out on glass and hydroxyapatite discs because there are surface-specific effects on S. mutans attachment and biofilm formation (16). S. mutans attachment on glass and hydroxyapatite was increased by 15 and 6 times, respectively, when sucrose was present compared to when glucose was present
(Figure 2.1A and C). S. mutans biofilm formation in the presence of sucrose was increased by
45% on glass and 8% on hydroxyapatite compared to that in the presence of glucose (Figure
2.1B and D).
30 Figure 2.1
31 Figure 2.1 (Continued). Sucrose enhances S. mutans attachment and biofilm formation. The levels of S. mutans attachment (A) and biofilm formation (B) on glass are significantly enhanced at all time points when the bacteria are grown in BHI containing 1% sucrose (Medium+Sucrose) compared to the levels achieved in BHI containing 1% glucose (Medium+Glucose). S. mutans attachment (C) and biofilm formation (D) on hydroxyapatite are similarly increased in the presence of sucrose, illustrating that the effect of sucrose on S. mutans physiology is not surface specific. Fluorescence microscopy images verify the findings of the CFU count experiments by showing an increase in S. mutans attachment (E) and biofilm formation (F) on glass in the presence of sucrose. *, statistically significant difference determined by Student’s t test (P<0.02). Error bars represent SDs. Scale bars, 20 µm.
The number of S. mutans cells in the biofilm for each condition is represented as a fraction of the total number of bacteria in the well, because S. mutans’ growth rate changes in the presence of sucrose compared with that in the presence of glucose (Figure 2.2).
Figure 2.2
Figure 2.2 Supplemental sugar alters S. mutans growth. A growth curve of S. mutans in BHI with added 1% glucose (Medium+Glucose) or 1% sucrose (Medium+Sucrose) shows that S. mutans’ growth rate changes based on the specific sugar present in the growth medium. Error bars represent SDs.
The increase in S. mutans attachment and biofilm formation in the presence of sucrose is supported by a fluorescence microscopy time series using SYTO9 nucleic acid stain to visualize
S. mutans attachment and biofilm formation in BHI containing 1% sucrose or 1% glucose on a
32 glass surface (Figure 2.1E and F). Our results support those of previous studies by quantitatively showing that the addition of 1% sucrose enhanced S. mutans attachment and biofilm formation at all time points compared to attachment and biofilm formation in the presence of 1% glucose (12–
15). There was little or no growth in BHI without a sugar source, illustrating that protein alone cannot support S. mutans attachment or biofilm formation. Furthermore, our results showing that sucrose enhances S. mutans attachment and biofilm formation were consistent on hydroxyapatite and glass surfaces, indicating that the effect is not surface specific. These findings set the groundwork for our investigation of the role of MUC5B in S. mutans attachment and biofilm formation. When testing the effect of MUC5B on S. mutans physiology, 1% sucrose was added to BHI medium to challenge the effect of MUC5B by ensuring that S. mutans attachment and biofilm formation are most robust.
MUC5B decreases S. mutans attachment and biofilm formation.
The effect of MUC5B on S. mutans attachment and biofilm formation was evaluated using CFU counts on various surfaces and fluorescence microscopy. S. mutans (107 bacteria) was grown in a chambered glass slide or on a hydroxyapatite disc in the presence of BHI with 1% sucrose and 0.3% MUC5B. BHI with 1% sucrose and 0.3% methylcellulose and BHI with no added polymer served as controls. Methylcellulose is a gel-forming compound that, like mucins, imparts viscosity but does not contain the complex, glycosylated structure that is characteristic of
MUC5B. On glass, the addition of MUC5B to growth medium decreased S. mutans attachment by 88% and biofilm formation by 74% compared to the levels of attachment and biofilm formation in BHI with 1% sucrose (Figure 2.3A and B). In comparison, the addition of methylcellulose reduced S. mutans attachment and biofilm formation on glass by 50% and 16%, respectively (Figure 2.3A and B). When S. mutans was grown on hydroxyapatite discs in the
33 presence of MUC5B, attachment was decreased by 77% and biofilm formation was decreased by
95% compared to the levels in BHI with 1% sucrose (Figure 2.3C and D). In comparison, the presence of methylcellulose reduced S. mutans attachment by 27% and biofilm formation by
76% on hydroxyapatite discs (Figure 2.3C and D). There was an overall decrease in biofilm formation at 18 and 24 h due to the dissolution of the hydroxyapatite discs. By evaluating the attachment and biofilm formation of S. mutans in medium containing methylcellulose, which simulates an environment that has physical properties similar to those of mucins, we can better understand if MUC5B is acting as a physical barrier to attachment most likely through increased viscosity or if the observed effect is due to specific MUC5B properties. We can conclude that the latter is most likely because S. mutans attachment and biofilm formation in the presence of
MUC5B are significantly decreased at most time points compared to the levels in the presence of methylcellulose. There are at least three potential mechanisms by which MUC5B could protect the surface from bacterial colonization: (i) MUC5B could bind or agglutinate bacteria, which would allow planktonic bacteria to be swept out of the oral cavity with salivary flow but enhance bacterial attachment to surfaces coated with MUC5B (17–26), (ii) MUC5B could have the opposite effect, where its heterogeneous glycan chains repel bacteria, thereby preventing surface attachment (4, 27–30), or (iii) MUC5B could directly downregulate S. mutans genes involved in attachment and biofilm formation. In our case, it appears that MUC5B is repelling S. mutans and/or directly influencing genetic modifications that protect the glass and hydroxyapatite surfaces from bacterial attachment and biofilm formation. The decrease in attachment caused by methylcellulose indicates that increased viscosity may also be playing some role in reducing attachment at early time points. Fluorescence microscopy experiments using SYTO9 staining confirm the findings obtained by CFU counts by showing a visually detectable decrease in the
34 amount of S. mutans on glass and that the characteristic microcolony morphology of S. mutans biofilms is unchanged (Figure 2.3E and F) (31, 32).
Figure 2.3
Figure 2.3 Salivary mucins reduce S. mutans attachment and biofilm formation. The addition of 0.3% mucins to the control medium, BHI containing 1% sucrose (SMedium), significantly reduces the levels of S. mutans attachment and biofilm formation on glass (A, B)
35 Figure 2.3 (Continued) and hydroxyapatite (C, D) compared to the levels obtained with the control consisting of BHI with 1% sucrose. Similarly, the addition of 0.3% methylcellulose to BHI with 1% sucrose reduces S. mutans attachment and biofilm formation; however, the effect is not significant for the majority of time points studied. Fluorescence microscopy was used to visually assess S. mutans attachment (E) and biofilm formation (F) on glass when the bacteria are grown in BHI with 1% sucrose and 0.3% mucins, BHI with 1% sucrose and 0.3% methylcellulose (Methyl.), and BHI with 1% sucrose. *, statistically significant difference from BHI with 1% sucrose determined by Student’s t test (P<0.02). Error bars represent SDs. Scale bars, 20 µm.
MUC5B does not alter S. mutans growth.
To evaluate the effect of MUC5B on bacterial growth, S. mutans was grown in BHI medium containing 1% sucrose and 0.3% MUC5B. BHI with 1% sucrose and 0.3% methylcellulose and BHI with 1% sucrose were used as controls. These are the same media used in experiments that determine the effects of MUC5B on S. mutans attachment and biofilm formation. Optical density readings over the course of 12 h show that the addition of MUC5B or methylcellulose does not alter S. mutans growth compared to its growth in BHI containing 1% sucrose (Figure 2.4). Because the growth of S. mutans is unchanged by MUC5B, we can conclude that the observed decrease in S. mutans attachment and biofilm formation in the presence of MUC5B is not due to slower growth but, rather, is due to the intrinsic properties of the MUC5B glycoprotein.
36 Figure 2.4
Figure 2.4 S. mutans growth is unaffected by the presence of salivary mucins. A growth curve of S. mutans in BHI with 1% sucrose (SMedium), BHI with 1% sucrose and 0.3% mucins, or BHI with 1% sucrose and 0.3% methylcellulose indicates that the presence of mucins and methylcellulose does not alter the growth of S. mutans. Error bars represent SDs.
MUC5B keeps S. mutans in planktonic form.
By quantifying the biofilm and supernatant bacteria in S. mutans cultures grown from 6 to 24 h, we determined the effect of MUC5B on S. mutans over time after stationary phase is reached. When the numbers of planktonic and biofilm S. mutans CFU are combined to determine the total number of viable S. mutans cells in a given experiment, results show that there is no significant difference between the total numbers of live bacteria in cultures containing 0.3%
MUC5B and in control media (Figure 2.5). Based on these findings, we show that the reduction in S. mutans attachment and biofilm formation on glass and hydroxyapatite in the presence of
MUC5B (Figure 2.3) is not due to bactericidal properties of MUC5B. The presence of MUC5B reduces S. mutans attachment and biofilm formation by maintaining bacteria in the planktonic phase. These findings point to the importance of MUC5B in establishing a healthy oral microbiota that allows species diversity but, at the same time, protects teeth from bacterial
37 damage.
Figure 2.5
Figure 2.5 S. mutans survival is unaffected by salivary mucins. The graph represents the total number of viable S. mutans cells per well in the supernatant and biofilm in BHI with 1% sucrose (SMedium), BHI with 1% sucrose and 0.3% mucins, or BHI with 1% sucrose and 0.3% methylcellulose. Salivary mucins and methylcellulose show no bactericidal effects at time points up to 24 h. Error bars represent SDs.
Conclusion
In summary, we used CFU counts and fluorescence light microscopy to quantitatively show that S. mutans attachment and biofilm formation are most robust when the organism is grown in the presence of sucrose and that the addition of purified human salivary MUC5B significantly decreases S. mutans attachment and biofilm formation even in the presence of sucrose. We determined that MUC5B does not alter S. mutans growth or lead to bacterial killing over 24 h but limits biofilm formation by maintaining S. mutans primarily in the planktonic form
(Figure 2.6). We speculate that the observed decrease in bacterial attachment and biofilm formation is due to a combination of genetic changes that decrease bacterial surface colonization and repulsion by MUC5B’s heterogeneous glycans. S. mutans attachment and biofilm formation
38 are key steps in the development of dental caries; therefore, these findings have particularly important clinical implications. The presence or absence of MUC5B in the oral cavity could alter individuals’ susceptibility to dental cavity formation, which could then be an easily accessible, highly predictable clinical diagnostic marker of disease. From a therapeutic standpoint, exogenous MUC5B could potentially be utilized as a treatment or preventative measure for dental caries. These findings illustrate that MUC5B may help protect teeth from cavity formation, but further studies, such as those that use RNA sequencing or other genetic profiling techniques, are needed to fully characterize the mechanism underlying the observed decrease in S. mutans attachment and biofilm formation.
Figure 2.6 ABmucins S. mutans
Figure 2.6 Summary of conclusions. S. mutans utilizes sucrose to form sticky extracellular polysaccharides that facilitate attachment to the tooth surface and subsequent biofilm formation. (A) In the biofilm, bacterial metabolism of sucrose causes a decrease in the local pH, leading to demineralization of the tooth structure. (B) The presence of mucins in sucrose-supplemented growth medium decreases S. mutans attachment and biofilm formation on the tooth surface by maintaining S. mutans in the planktonic state. Mucin illustrations represent multimers. Figure not drawn to scale.
39 Materials and Methods
Bacterial strains and growth conditions. The bacterial strain Streptococcus mutans UA159 was kindly given as a gift by Dan Smith (Forsyth Institute). For sucrose and glucose experiments,
S. mutans was grown overnight in brain heart infusion (BHI) medium (Becton, Dickinson and
Company) containing 1% (wt/vol) sucrose and BHI with 1% (wt/vol) glucose (Sigma). For experiments determining the effects of MUC5B, S. mutans was grown overnight in BHI with 1% sucrose. BHI with 1% sucrose and either 0.3% MUC5B or 0.3% (wt/vol) methylcellulose
(Sigma) was used to resuspend S. mutans cells before inoculating them into the experiment.
Hydroxyapatite disc (Clarkson Chromatography, Inc.) or chambered glass slide (LabTek) surfaces were used to test S. mutans attachment and biofilm formation. S. mutans was grown and incubated at 37°C with 5% CO2.
Saliva collection. Submandibular saliva was collected from 10 volunteers using a custom vacuum pump setup. Specifically, two holes were cut into the cap of a 50-ml conical tube
(Falcon); the vacuum line was inserted into one hole and a small-diameter Tygon collection tube was inserted into the other hole (Saint Gobain Performance Plastics). Cotton swabs were used to absorb the volunteers’ parotid gland secretions. The collection tube was used to suck up pooled unstimulated submandibular gland secretions from under the tongue. The collection vessel was kept on ice at all times. Saliva from volunteers was pooled before MUC5B purification.
Protocols involving the use of human subjects were approved by Massachusetts Institute of
Technology’s Committee on the Use of Humans as Experimental Subjects.
MUC5B purification. Immediately after collection, saliva was diluted using 5.5 M sodium chloride containing 0.04% sodium azide so that the final concentration of sodium chloride was
0.16 M. The following antibacterial agents and protease inhibitors were then added at the
40 indicated final concentrations: benzamidine HCl (5 mM), dibromoacetophenone (1 mM), phenylmethylsulfonyl fluoride (1 mM), and EDTA (5 mM, pH 7) (Sigma). The mucins in the saliva were solubilized overnight by gentle stirring at 4°C. Saliva was then centrifuged at 3,800 g for 10 min in a swinging-bucket centrifuge to remove cellular debris. MUC5B was purified using a Bio-Rad NGC fast protein liquid chromatography (FPLC) system equipped with an XK 50 column packed with Sepharose CL-2B resin (GE Healthcare Bio-Sciences). Mucin-containing fractions were identified using a periodic acid-Schiff’s reagent assay and analysis of UV absorbance at 280 nm from FPLC. Fractions were then combined, dialyzed, and concentrated using an ultrafiltration device and were then lyophilized for storage at 80°C.
Assay of CFU counts to evaluate S. mutans attachment and biofilm formation. To test the effects of sucrose or glucose on S. mutans physiology, S. mutans was grown to mid-exponential phase in BHI with 1% sucrose and BHI with 1% glucose, and then equal numbers of bacteria
(107) from each culture were seeded in triplicate into wells containing glass or hydroxyapatite surfaces. For experiments testing the effect of MUC5B, S. mutans was grown to mid-exponential phase in BHI with 1% sucrose and then seeded in triplicate into wells containing BHI with 1% sucrose and 0.3% MUC5B or control medium. For all experiments, attachment was evaluated at
20, 40, and 60 min and biofilm formation was evaluated at 6, 18, and 24 h. Attachment was defined to occur at time points up to 1 h because the doubling time of S. mutans is approximately
1.5 h. Biofilm formation was defined to occur at all time points after 1 h. At the time point being evaluated, the surface was washed with phosphate buffered saline (PBS) to remove nonadherent cells, fresh PBS was added, and then adherent cells were lifted using a sterile pipette tip. The suspended bacteria were vigorously pipetted to individualize the cells. The suspension was diluted (10-1 to 10-7) and plated on BHI agar. The numbers of CFU were counted after 24 to 36 h
41 of incubation. Statistically significant differences were determined using Student’s t test, with P values of <0.02 considered significant.
Fluorescent staining and microscopy. To visually assess the effects of sucrose, glucose, or
MUC5B on S. mutans attachment and biofilm formation, fluorescent SYTO9 staining with light microscopy was used (Life Technologies). S. mutans was grown to mid-exponential phase, then seeded into a chambered glass slide with BHI containing 1% sucrose and 0.3% MUC5B or with control medium. At the time point being evaluated, the surface was washed with PBS to remove nonadherent cells, and then 200 µl SYTO9 (0.6 µl SYTO9/200 µl Milli-Q water) was added. The biofilm was incubated with SYTO9 in the dark for 30 min. After incubation, the biofilm was washed with Milli-Q water to remove excess dye and fresh Milli-Q water was added. A Zeiss
Axio Observer Z1 fluorescence inverted microscope was used for imaging. All experiments were repeated in triplicate.
Assay to evaluate S. mutans growth. Overnight cultures of S. mutans at an optical density at
600 nm (OD600) of 0.05 were seeded in triplicate into a 96-well polystyrene plate containing BHI medium supplemented with 1% sucrose and 0.3% MUC5B or control medium. Bacteria were incubated at 37°C with 5% CO2. At 1-h intervals, the cultures were mixed and the OD600 was recorded using a microplate reader. The averages for each time point were plotted, and a comparison of S. mutans’ growth rates in the various media was evaluated within the estimated error.
Time-kill assay. CFU counts were used to evaluate the effect of MUC5B on S. mutans viability at time points up to 24 h. S. mutans was grown to mid-exponential phase in BHI with 1% sucrose, and then equal numbers of bacteria (107) were seeded in triplicate into glass-bottom wells
42 containing BHI with 1% sucrose and 0.3% MUC5B or control medium. The cultures were incubated for 6, 18, and 24 h. At the time point being evaluated, the contents of the wells were gently mixed and then the supernatant was removed and diluted (10-1 to 10-8). The remaining biofilm was then washed with PBS, scraped off with a sterile pipette tip, and diluted (10-1 to 10-6).
Dilutions were plated on BHI agar. The numbers of CFU were counted after 24 to 36 h of incubation to quantify the number of viable bacteria.
Acknowledgements
We thank Nicole Kavanaugh for her critical review of the figures.
This work was made possible by generous funding support from Colgate-Palmolive (to K.R.) and Biological Sciences in Dental Medicine grants for tuition and a stipend (to E.S.F.).
43 References
1. Brockhausen I, Schachter H, Stanley P. 2009, posting date. Chapter 9, O-GalNAc Glycans. In Varki A, Cummings R, Esko J, Freeze H, Stanley P, Bertozzi C, Hart G, Etzler M (ed), Essentials of Glycobiology, 2nd ed, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
2. Perez-Vilar J, Mabolo R. 2007. Gel-forming mucins. Notions from in vitro studies. Histol Histopathol 22:455–464.
3. Tabak LA. 1995. In Defense of the Oral Cavity: Structure, Biosynthesis, and Function of Salivary Mucins. Annu Rev Physiol 57:547–564.
4. Loesche WJ. 1986. Role of Streptococcus mutans in human dental decay. Microbiol Rev 50:353–380.
5. Al-Hashimi I, Levine MJ. 1989. Characterization of in vivo salivary-derived enamel pellicle. Arch Oral Biol 34:289–295.
6. Cárdenas M, Elofsson U, Lindh L. 2007. Salivary Mucin MUC5B Could Be an Important Component of in Vitro Pellicles of Human Saliva: An in Situ Ellipsometry and Atomic Force Microscopy Study. Biomacromolecules 8:1149–1156.
7. Bowen WH, Koo H. 2011. Biology of Streptococcus mutans-derived glucosyltransferases: role in extracellular matrix formation of cariogenic biofilms. Caries Res 45:69–86.
8. Nakano YJ, Kuramitsu HK. 1992. Mechanism of Streptococcus mutans glucosyltransferases: hybrid-enzyme analysis. J Bacteriol 174:5639–5646.
9. Krzyściak W, Jurczak A, Kościelniak D, Bystrowska B, Skalniak A. 2014. The virulence of Streptococcus mutans and the ability to form biofilms. Eur J Clin Microbiol Infect Dis 33:499–515.
10. Cross SE, Kreth J, Zhu L, Sullivan R, Shi W, Qi F, Gimzewski JK. 2007. Nanomechanical properties of glucans and associated cell-surface adhesion of Streptococcus mutans probed by atomic force microscopy under in situ conditions. Microbiology 153:3124–3132.
11. Socransky SS, Haffajee AD. 2002. Dental biofilms: difficult therapeutic targets. Periodontol 2000 28:12–55.
12. Shemesh M, Tam A, Steinberg D. 2007. Expression of biofilm-associated genes of Streptococcus mutans in response to glucose and sucrose. J Med Microbiol 56:1528–1535.
13. Koo H, Xiao J, Klein MI, Jeon JG. 2010. Exopolysaccharides produced by Streptococcus mutans glucosyltransferases modulate the establishment of microcolonies within multispecies biofilms. J Bacteriol 192:3024–3032.
44 14. Duarte S, Klein MI, Aires CP, Cury JA, Bowen WH, Koo H. 2008. Influences of starch and sucrose on Streptococcus mutans biofilms. Oral Microbiol Immunol 23:206–212.
15. Kreth J, Zhu L, Merritt J, Shi W, Qi F. 2008. Role of sucrose in the fitness of Streptococcus mutans. Oral Microbiol Immunol 23:213–219.
16. Shemesh M, Tam A, Aharoni R, Steinberg D. 2010. Genetic adaptation of Streptococcus mutans during biofilm formation on different types of surfaces. BMC Microbiol 10:51.
17. Gibbons RJ, Cohen L, Hay DI. 1986. Strains of Streptococcus mutans and Streptococcus sobrinus attach to different pellicle receptors. Infect Immun 52:555–561.
18. Tabak LA, Levine MJ, Mandel ID, Ellison SA. 1982. Role of salivary mucins in the protection of the oral cavity. J Oral Pathol 11:1–17.
19. McBride BC, Gisslow MT. 1977. Role of sialic acid in saliva-induced aggregation of Streptococcus sanguis. Infect Immun 18:35–40.
20. Houte PJ van. 1982. Bacterial adherence and dental plaque formation. Infection 10:252–260.
21. Levine MJ, Herzberg MC, Levine MS, Ellison SA, Stinson MW, Li HC, van Dyke T. 1978. Specificity of salivary-bacterial interactions: role of terminal sialic acid residues in the interaction of salivary glycoproteins with Streptococcus sanguis and Streptococcus mutans. Infect Immun 19:107–115.
22. Klein A, Carnoy C, Wieruszeski JM, Strecker G, Strang AM, Van Halbeek H, Roussel P, Lamblin G. 1992. The broad diversity of neutral and sialylated oligosaccharides derived from human salivary mucins. Biochemistry 31:6152–6165.
23. Williams RC, Gibbons RJ. 1975. Inhibition of streptococcal attachment to receptors on human buccal epithelial cells by antigenically similar salivary glycoproteins. Infect Immun 11:711–718.
24. Thomsson KA, Prakobphol A, Leffler H, Reddy MS, Levine MJ, Fisher SJ, Hansson GC. 2002. The salivary mucin MG1 (MUC5B) carries a repertoire of unique oligosaccharides that is large and diverse. Glycobiology 12:1–14.
25. Gibbons RJ, Dankers I. 1981. Lectin-Like Constituents of Foods Which React with Components of Serum, Saliva, and Streptococcus mutans. Appl Environ Microbiol 41:880– 888.
26. Gibbons RJ, Qureshi JV. 1978. Selective binding of blood group-reactive salivary mucins by Streptococcus mutans and other oral organisms. Infect Immun 22:665–671.
45 27. Veerman EC, Ligtenberg AJ, Schenkels LC, Walgreen-Weterings E, Nieuw Amerongen AV. 1995. Binding of human high-molecular-weight salivary mucins (MG1) to Hemophilus parainfluenzae. J Dent Res 74:351–357.
28. Murray PA, Prakobphol A, Lee T, Hoover CI, Fisher SJ. 1992. Adherence of oral streptococci to salivary glycoproteins. Infect Immun 60:31–38.
29. Amerongen AVN, Bolscher JGM, Veerman ECI. 1995. Salivary mucins: protective functions in relation to their diversity. Glycobiology 5:733–740.
30. Schenkels LCPM, Ligtenberg AJM, Veerman ECI, Van Nieum Amerongen A. 1993. Interaction of the Salivary Glycoprotein EP-GP with the Bacterium Streptococcus salivarius HB. J Dent Res 72:1559–1565.
31. Xiao J, Koo H. 2010. Structural organization and dynamics of exopolysaccharide matrix and microcolonies formation by Streptococcus mutans in biofilms. J Appl Microbiol 108:2103– 2113.
32. Xiao J, Klein MI, Falsetta ML, Lu B, Delahunty CM, Yates JR III, Heydorn A, Koo H. 2012. The Exopolysaccharide Matrix Modulates the Interaction between 3D Architecture and Virulence of a Mixed-Species Oral Biofilm. PLoS Pathog 8:e1002623.
46 Chapter III
Salivary Mucins Promote the Coexistence of Competing Oral Bacterial
Species
Results presented in this chapter are in press:
Frenkel ES, Ribbeck K. Salivary mucins promote the coexistence of competing oral bacterial
species. ISME J.
47 Abstract
Mucus forms a major ecological niche for microbiota in various locations throughout the human body, such as the gastrointestinal tract, respiratory tract and oral cavity. The primary structural components of mucus are mucin glycoproteins, which crosslink to form a complex polymer network that surrounds microbes. Although the mucin matrix could create constraints that impact inhabiting microbes, little is understood about how this key environmental factor affects interspecies interactions. In this study, we develop an experimental model using gel- forming human salivary mucins to understand the influence of mucin on the viability of two competing species of oral bacteria. We use this dual-species model to show that mucins promote the coexistence of the two competing bacteria and that mucins shift cells from the mixed-species biofilm into the planktonic form. Taken together, these findings indicate that the mucus environment could influence bacterial viability by promoting a less competitive mode of growth.
Introduction
Mucus lines wet epithelia throughout the human body and is a major ecological niche for microbiota in the respiratory tract, gastrointestinal tract and oral cavity among other locations (1–
3). The mucus layer is a three dimensional hydrogel primarily composed of densely glycosylated polymers called mucins (4, 5). In the gel, mucin chains cross-link to form a network that surrounds microbes and, consequently, could create geometric and diffusive constraints for biotic and abiotic environmental factors. Little is known, however, about how these constraints influence microbial interactions, such as cell-cell communication and competition, among the vast number of organisms that live in mucus. In this study, we build upon our previous work
48 showing that MUC5B mucins affect intraspecies interactions by promoting dispersal of bacteria and fungi (6–8). Here, we develop an experimental model to probe the influence of gel-forming human salivary mucins on dual-species bacterial competition to understand how this matrix affects the viability of competing bacteria.
Results and Discussion
The dual-species model is composed of Streptococcus sanguinis JFP36 and S. mutans
UA159, which compete in the oral cavity through the production of hydrogen peroxide and antimicrobial peptides called mutacins, respectively (9–12). In this model, the two species were inoculated sequentially to more closely mimic the natural environment of the oral cavity where surfaces are generally coated by microbes before other species attempt to colonize. When S. mutans was the primary colonizer, and MUC5B mucins were not present in the growth medium, viability of the secondary colonizer (S. sanguinis) rapidly declined, suggesting that S. mutans outcompetes S. sanguinis (Fig 3.1 A(I), (II)). In contrast, when MUC5B was present in the growth medium, the total number of viable S. sanguinis cells increased by 18- and 88-fold after 4 and 5 h of co-culture, respectively, compared to the control without mucin (Figure 3.1 A(II)).
The same protective effect by MUC5B was observed when S. sanguinis was the primary colonizer. In this case, the addition of MUC5B to medium enhanced survival of both S. sanguinis and S. mutans (Figure 3.1 B(I), (II)). After 4 and 5 h of co-culture in the presence of MUC5B, the number of viable S. sanguinis cells increased by 9- and 94-fold, respectively, relative to the control (Figure 3.1 B(I)). S. mutans CFU increased by 2-, 3-, and 7-fold at 4, 5 and 6 h, respectively (Figure 3.1 B(II)).
49 Figure 3.1 Figure 1
A (I). SMedium SMedium+Mucin A (II). 108 108
6 6 10 10 * * 104 104 S. mutans S. sanguinis 102 102 Total Total
0 Total 0
cell population (CFU) 10 10 Start 3 4 5 cell population (CFU) Start 3 4 5 Time of co-culture (h) Time of co-culture (h) B (I). B (II). 1010 108 8 * 10 106 * 106 * 104 4 * 10 S. mutans S. sanguinis * 2 102 10 Total Total Total Total 100 100 cell population (CFU) Start 4 5 6 cell population (CFU) Start 4 5 6 Time of co-culture (h) Time of co-culture (h) s C. SMedium SMedium+Mucin D. s Primary colonizer: S. sanguinis 105 Primary colonizer: S. mutans 105 4 sanguini 4 sanguini 10 . 10 . 3 S S
10 / 103 / 2 s ReducedReduced s 10 diversity 1 an 2 coexistence an 10 t 10 t 100 mu 1 mu 10 -1 . . 10 S S
100 10-2 io io t Start 3 4 5 t Start 4 5 Ra Time of co-culture (h) Ra Time of co-culture (h)
Figure 3.1 MUC5B promotes S. mutans and S. sanguinis coexistence. S. mutans and S. sanguinis viability in a dual-species experimental model containing control medium (half- strength BHI with 1% sucrose; SMedium) and control medium containing 0.4% MUC5B mucin. Viability was studied when S. mutans was the primary colonizer (A (I)) and S. sanguinis was the secondary colonizer (A (II)) and the reverse scenario where S. sanguinis was the primary colonizer (B (I)) and S. mutans was the secondary colonizer (B (II)). (C, D) The ratios of viable S. mutans and S. sanguinis when S. mutans was the primary colonizer (C) and when S. sanguinis was the primary colonizer (D) as an indication of species coexistence. *, statistically significant increase relative to the control with half-strength BHI containing 1% sucrose determined by Student’s t test (P<0.05). Experiments were performed in triplicate and error bars represent SD of CFU between replicates.
50 Of note is that although MUC5B significantly enhanced S. sanguinis viability, there was an overall reduction in S. sanguinis CFU due to self-killing, which could be caused by increasing hydrogen peroxide concentrations. The same reduction in viability was observed when S. sanguinis was grown in monoculture, indicating that the killing was not due to S. mutans (Figure
3.2).
Figure 3.2
Supplemental Figure 1 )
10
U 10 s F 8
(C 10
6
ion 10 t sanguini . 104 * S
al 2 t popula 10
o * T 100 * cell Start 7 8 9 Time (h)
Figure 3.2 At high cell density, S. sanguinis viability decreases over time due to self-killing. When S. sanguinis is grown at high cell densities in a mono-species culture, the number of viable cells decreases over 9 hours. *, statistically significant decrease relative to the starting cell population determined by Student’s t test (P<0.05). Experiments were performed in triplicate and error bars represent SD of CFU between replicates.
Methylcellulose, a gel-forming polymer that is commonly used to mimic the viscosity of mucus, did not have the same protective effect on S. mutans or S. sanguinis in both of these experimental models; the enhanced survival is likely not due to the addition of a polymer, which could increase viscosity or cause osmotic stress (Figure 3.3) (13, 14). Together these results show that
MUC5B significantly enhances bacterial coexistence by increasing survival of at least one bacterial population compared to the control without MUC5B (Figure 3.1 C, D).
51 Figure 3.3 Supplemental Figure 2 SMedium SMedium+Methyl. ) )
8 8 U U
A (I). 10 A (II). s 10 F F s (C (C an
6 6 t 10 10 ion ion mu t t sanguini . 104 104 . S
S
al
t 2 2 al
10 t 10 popula popula o
o T 0 T 0 cell 10 cell 10 Start 3 4 5 Start 3 4 5 Time of co-culture (h) Time of co-culture (h) ) B (I). B (II). ) 10 8 U U
s 10 10 F F s 8 (C (C an
10 6 t 10 6 ion ion mu t 10 t sanguini . 104 .
4 S
S 10
al
t 2 al
t 2 10 popula popula o
10 o T T 0 0 cell 10 cell 10 Start 4 5 6 Start 4 5 6 Time of co-culture (h) Time of co-culture (h)
Figure 3.3. Methylcellulose does not affect S. mutans and S. sanguinis coexistence. S. mutans and S. sanguinis viability in a dual-species experimental model containing control medium (half- strength BHI with 1% sucrose; SMedium) and control medium containing 0.4% methylcellulose (Methyl.). Viability was studied when S. mutans was the primary colonizer (A (I)) and S. sanguinis was the secondary colonizer (A (II)) and the reverse scenario where S. sanguinis was the primary colonizer (B (I)) and S. mutans was the secondary colonizer (B (II)). Experiments were performed in triplicate and error bars represent SD of CFU between replicates.
To better understand how MUC5B influences bacterial viability, we studied several aspects of S. mutans and S. sanguinis growth in the presence of MUC5B. First, we determined that the observed protective effect of MUC5B was not due to increased bacterial growth;
MUC5B slightly reduces or has no effect on S. mutans and S. sanguinis growth rates (Figure 3.4
A, B). In addition, S. mutans and S. sanguinis did not grow in PBS containing mucin indicating that MUC5B is not used as a nutrient source under the conditions studied (Figure 3.4 A, B).
52 Another way MUC5B could increase bacterial survival is by altering the cells’ mode of growth, which can influence interspecies competition and cell properties such as gene regulation and cell-cell communication (15–20). Our data show that MUC5B efficiently reduces S. mutans and
S. sanguinis biofilm formation in single-species cultures at 6 h and 9 h: S. mutans biofilm formation was reduced by 17- and 12-fold and S. sanguinis biofilm formation decreased 16- and
8-fold compared to the control without mucin (Figure 3.4 C(I) and (II), D (I) and (II)). Strikingly, the total S. mutans and S. sanguinis cell populations were unchanged at all time points in the presence of MUC5B (except for a slight decrease at 6 h, but the number of cells in the biofilm at this time still account for only 13% of total cells) (Figure 3.4 C(I) and (II), D (I) and (II)). This result implies that, in the presence of MUC5B, the vast majority of cells shifted into the planktonic state. Because the single cell bacterial form can be less competitive than the surface- attached state, this movement of cells away from the biofilm could be a mechanism to reduce interspecies competition. After 4 h of co-culture in the dual-species model, we found that
MUC5B also reduced biofilm formation of the primary colonizer in the mixed biofilm by 19-fold for S. mutans and 3-fold for S. sanguinis relative to the control without polymer (Figure 3.4 E,
F). As shown in Figure 3.1 A(I) and B(I), the total cell population in each of these cases was unaffected (S. mutans) or increased (S. sanguinis) by MUC5B, indicating a decrease in the relative proportion of biofilm cells. In the case of the secondary colonizer, there was also an overall reduction in the proportion of biofilm cells for both S. mutans and S. sanguinis; the total number of viable cells increased in the presence of MUC5B (Figure 3.1 A(II), B(II)), yet there was only a slight increase or no change in biofilm formation in the presence of MUC5B relative to the control without polymer (Figure 3.4 E, F). Taken together, these results indicate that
MUC5B could enhance bacterial coexistence and possibly bacterial diversity in the oral cavity
53 by shifting competing species away from the biofilm and into the less competitive planktonic state, although further studies would be needed to establish this effect.
54 Figure 3.4
Figure 2 SMedium PBS+Mucin A. SMedium+Mucin B.
8 ) 8
) 10 10 U F (CFU
(C 7 7
10 s 10 ans t 106 106 mu sanguini . . S 105 S 105 0 2 4 6 8 10 0 2 4 6 8 10
Time (h) Time (h)
C (I). Total cell population Biofilm population C (II).
8 6 h 8 9 h 10 ion 10 t ) ) 7 7 10 U 10 U * F popula F
population s (C
(C
ans an 6 6 t t 10 * 10 * mu mu
. . 5 5 S S 10 10 SMedium SMedium+ SMedium SMedium+ Mucin
D (I). D (II). Mucin ion ion 8 8 t t 10 6 h 10 9 h 7 7
10 ) 10 ) U U popula popula
6 F 6
F 10 10 (C
(C 105 105 104 * 104 *
sanguinis sanguinis . . 103 103 S S SMedium SMedium+ SMedium SMedium+ Mucin Mucin ) ) U E. F. U F F 8 8 (C Primary colonizer: S. mutans (C Primary colonizer: S. sanguinis
10 10 S. mutans ion ion t t 106 6 * 10 S. mutans popula popula
104 104 ilm ilm f S. sanguinis f S. sanguinis
Bio 2 10 Bio 102 * SMedium SMedium+ SMedium SMedium+ Mucin Mucin -mucin +mucin G. S. mutans / S. sanguinis H. mucin
SMe-dium
55 Figure 3.4 (Continued) MUC5B reduces S. mutans and S. sanguinis surface attachment by shifting cells into the planktonic state. S. mutans (A) and S. sanguinis (B) growth in control medium (half-strength BHI with 1% sucrose; SMedium), control medium containing 0.4% MUC5B mucin, and PBS containing 0.4% MUC5B mucin. (C (I), (II)) S. mutans biofilm population and total cell population in mono-species cultures containing control medium and control medium with 0.4% MUC5B mucin at 6 h (C (I)) and 9 h (C (II)). (D (I), (II)) S. sanguinis biofilm population and total cell population in mono-species cultures containing control medium and control medium with 0.4% MUC5B mucin at 6 h (D (I)) and 9 h (D (II)). (E, F) S. mutans and S. sanguinis biofilm formation in control medium and control medium containing 0.4 % MUC5B mucin in a dual-species model after 4 h of co-culture when S. mutans was the primary colonizer (E) and when S. sanguinis was the primary colonizer (F). (G, H) Schematic illustrating a summary of conclusions. In the absence of MUC5B (G), biofilm formation of S. mutans and S. sanguinis increases and bacterial coexistence decreases compared to biofilm formation and species coexistence in the presence of mucin (H). Mucin illustrations represent multimers. Schematic not drawn to scale. *, statistically significant decrease relative to the control with half- strength BHI containing 1% sucrose determined by Student’s t test (P<0.05). Experiments were performed in triplicate and error bars represent SD of CFU between replicates.
Conclusion
In this work, we use a dual-species bacterial model containing human MUC5B salivary mucin to understand how this prevalent environmental factor influences bacterial viability. Our results show that: 1) MUC5B promotes S. mutans and S. sanguinis coexistence, and 2) MUC5B shifts cells from the biofilm into the planktonic state (Figure 3.4 G, H). By promoting the single cell (planktonic) state, MUC5B could alter cell-cell interactions, toxin production, or other mechanisms of competition. Although this model is not as complex as the oral cavity microbiota, these findings are among the first to indicate that mucus and its primary structural component, mucins, could influence bacterial survival in a multispecies environment. Further studies are needed, however, to understand if the observed increase in bacterial survival and reduction in surface colonization in the presence of mucin are due to an indirect influence of mucin, such as altered transport of secreted factors, or a direct impact on bacterial physiology, which could change gene regulation.
56 Material and Methods
Bacterial strains and growth conditions. Streptococcus mutans UA159 was kindly given as a gift by Dr. Dan Smith (Forsyth Institute) and erythromycin-resistant Streptococcus sanguinis
JFP36 was generously given by Dr. Todd Kitten (Virginia Commonwealth University) (9, 12).
For all experiments, S. mutans and S. sanguinis were grown overnight in Brain Heart Infusion
(BHI) medium (Becton, Dickinson and Company). Before inoculating an experiment, the cells were pelleted, washed with half-strength BHI containing 1% (wt/vol) sucrose (Sigma), then resuspended in half-strength BHI containing 1% sucrose. Acid-washed glass beads (425-600 µm,
Sigma) were added to the resuspended bacteria to break up bacterial aggregates then the culture was vortexed for ten pulses of two seconds each. To determine the effect of methylcellulose and
MUC5B, bacteria were resuspended in BHI with 1% sucrose and either 0.4% (wt/vol) MUC5B or 0.4% methylcellulose (Sigma) before inoculating them into an experiment. All experiments were performed in chambered glass slides (LabTek) at 37°C with 5% CO2.
Saliva collection. Submandibular saliva was collected from 9 volunteers using a custom vacuum pump as previously described (7). Briefly, a vacuum line and a collection tube (Saint Gobain
Performance Plastics) were inserted into a conical tube (Falcon). Unstimulated submandibular gland secretions were collected from under the tongue and the collection vessel was kept on ice at all times. Saliva from individual volunteers was combined prior to purification. Protocols involving the use of human subjects were approved by Massachusetts Institute of Technology’s
Committee on the Use of Humans as Experimental Subjects.
MUC5B purification. MUC5B was purified as previously described (7). Briefly, saliva was diluted using sodium chloride containing sodium azide then antibacterial agents and protease
57 inhibitors were added. The saliva was stirred overnight at 4°C then centrifuged at 10,000 g for 10 min in a fixed-angle centrifuge to remove cellular debris. A Bio-Rad NGC fast protein liquid chromatography (FPLC) system with a Sepharose CL-2B column (GE Healthcare Bio-Sciences) were used to purify MUC5B. Mucin-containing fractions were then dialyzed and concentrated using an ultrafiltration device. Samples were lyophilized overnight for storage at -80°C. An enzyme-linked immunosorbent assay (ELISA) confirmed that the amount of MUC7 in the purified MUC5B is negligible; in a solution of 0.4% (w/v) MUC5B, MUC7 accounted for
0.0002% of the mucin present (LifeSpan BioSciences, Inc.).
CFU count assay to quantify total cell populations in dual-species models. S. mutans and S. sanguinis were suspended in half-strength BHI with 1% sucrose, half-strength BHI with 1% sucrose and 0.4% methylcellulose, or half-strength BHI with 1% sucrose and 0.4% MUC5B. For dual-species models where S. mutans was the primary colonizer, S. mutans (107) was inoculated and incubated for 3 hours then S. sanguinis (106) was inoculated. For dual-species models where
S. sanguinis was the primary colonizer, S. sanguinis (108) was inoculated, incubated for 3 hours, then S. mutans (107) was inoculated. After the secondary colonizer was inoculated, the dual- species culture was incubated until the time points indicated. At the end of the experiment a sterile pipette tip was used to scrape the biofilm, then the suspension of biofilm and supernatant cells was vortexed with glass beads. Cell suspensions were diluted then plated on BHI agar containing 10 µg/ml erythromycin to select for S. sanguinis and BHI agar with 1 U/ml bacitracin to select for S. mutans. The addition of antibiotics to agar was determined to have no significant bactericidal effect. Agar was incubated for 24-36 hours. The effect of methylcellulose and
MUC5B were determined relative to the control without added MUC5B or methylcellulose.
58 Statistically significant differences were determined using the Student’s t test, with P<0.05 considered significant.
S. mutans and S. sanguinis growth curve. Overnight cultures of S. mutans and S. sanguinis were resuspended in half-strength BHI with 1% sucrose, half-strength BHI with 1% sucrose and
0.4% MUC5B, or PBS with 0.4% MUC5B. The suspensions were vortexed with glass beads to individualize cells, then approximately 106 cells were inoculated into wells of a 96-well polystyrene plate. At 2-hour intervals the bottom of the wells were scraped to remove any adherent cells, mixed, and an aliquot was removed, diluted and plated on BHI agar. Agar was incubated and CFU were counted after 24-36 hours.
CFU count assay to quantify biofilm and total cell populations in single-species models.
S. mutans and S. sanguinis were suspended in half-strength BHI with 1% sucrose or half-strength
BHI with 1% sucrose and 0.4% MUC5B. Equal numbers of CFU from each species were inoculated into chambered glass slides then incubated. After a given amount of time, the supernatant was used to gently wash the biofilm to resuspend unattached cells, then the supernatant was removed. PBS was added to the wells then the biofilm was scraped using a sterile pipette tip. The supernatant and biofilm were vortexed with glass beads to individualize cells then the suspension was diluted, plated on BHI agar and incubated for 24-36 hours. The effect of MUC5B was determined relative to the control without an added polymer. Statistically significant differences were determined using the Student’s t test, with P<0.05 considered significant.
CFU count assay to quantify biofilm cell populations in dual-species models. S. mutans and
S. sanguinis were suspended in half-strength BHI with 1% sucrose or half-strength BHI with 1%
59 sucrose and 0.4% MUC5B. For dual-species models where S. mutans was the primary colonizer,
S. mutans (107) was inoculated and incubated for 3 hours then S. sanguinis (106) was inoculated.
For dual-species models where S. sanguinis was the primary colonizer, S. sanguinis (108) was inoculated, incubated for 3 hours, then S. mutans (107) was inoculated. After the secondary colonizer was inoculated, the dual-species culture was incubated for 4 h. After 4 h of co-culture the supernatant was used to gently wash the biofilm to resuspend unattached cells, then the supernatant was removed. PBS was added to the wells then the biofilm was scraped using a sterile pipette tip. Cell suspensions were diluted then plated on BHI agar containing 10 µg/ml erythromycin to select for S. sanguinis and BHI agar with 1 U/ml bacitracin to select for S. mutans. The addition of antibiotics to agar was determined to have no significant bactericidal effect. Agar was incubated for 24-36 hours. The effect of MUC5B was determined relative to the control without MUC5B. Statistically significant differences were determined using the
Student’s t test, with P<0.05 considered significant.
Acknowledgements
The authors would like to thank Julia Co, Nicole Kavanaugh and Gerardo Cárcamo-Oyarce for their helpful advice and guidance.
60 References
1. Derrien M, van Passel MW, van de Bovenkamp JH, Schipper RG, de Vos WM, Dekker J. 2010. Mucin-bacterial interactions in the human oral cavity and digestive tract. Gut Microbes 1:254–268.
2. Tabak LA. 1995. In Defense of the Oral Cavity: Structure, Biosynthesis, and Function of Salivary Mucins. Annu Rev Physiol 57:547–564.
3. Worlitzsch D, Tarran R, Ulrich M, Schwab U, Cekici A, Meyer KC, Birrer P, Bellon G, Berger J, Weiss T, Botzenhart K, Yankaskas JR, Randell S, Boucher RC, Döring G. 2002. Effects of reduced mucus oxygen concentration in airway Pseudomonas infections of cystic fibrosis patients. J Clin Invest 109:317–325.
4. Bansil R, Turner BS. 2006. Mucin structure, aggregation, physiological functions and biomedical applications. Curr Opin Colloid Interface Sci 11:164–170.
5. Tabak LA, Levine MJ, Mandel ID, Ellison SA. 1982. Role of salivary mucins in the protection of the oral cavity. J Oral Pathol 11:1–17.
6. Caldara M, Friedlander RS, Kavanaugh NL, Aizenberg J, Foster KR, Ribbeck K. 2012. Mucin Biopolymers Prevent Bacterial Aggregation by Retaining Cells in the Free- Swimming State. Curr Biol 22:2325–2330.
7. Frenkel ES, Ribbeck K. 2015. Salivary Mucins Protect Surfaces from Colonization by Cariogenic Bacteria. Appl Environ Microbiol 81:332–338.
8. Kavanaugh NL, Zhang AQ, Nobile CJ, Johnson AD, Ribbeck K. 2014. Mucins Suppress Virulence Traits of Candida albicans. mBio 5:e01911-14.
9. Ge X, Kitten T, Chen Z, Lee SP, Munro CL, Xu P. 2008. Identification of Streptococcus sanguinis Genes Required for Biofilm Formation and Examination of Their Role in Endocarditis Virulence. Infect Immun 76:2551–2559.
10. Kreth J, Zhang Y, Herzberg MC. 2008. Streptococcal Antagonism in Oral Biofilms: Streptococcus sanguinis and Streptococcus gordonii Interference with Streptococcus mutans. J Bacteriol 190:4632–4640.
11. Kreth J, Merritt J, Shi W, Qi F. 2005. Competition and Coexistence between Streptococcus mutans and Streptococcus sanguinis in the Dental Biofilm. J Bacteriol 187:7193–7203.
12. Senty Turner L, Das S, Kanamoto T, Munro CL, Kitten T. 2009. Development of genetic tools for in vivo virulence analysis of Streptococcus sanguinis. Microbiology 155:2573– 2582.
61 13. Ivic A, Onyeaka H, Girling A, Brewis IA, Ola B, Hammadieh N, Papaioannou S, Barratt CLR. 2002. Critical evaluation of methylcellulose as an alternative medium in sperm migration tests. Hum Reprod 17:143–149.
14. Smith DJ, Gaffney EA, Gadêlha H, Kapur N, Kirkman-Brown JC. 2009. Bend propagation in the flagella of migrating human sperm, and its modulation by viscosity. Cell Motil Cytoskeleton 66:220–236.
15. Kearns DB, Chu F, Branda SS, Kolter R, Losick R. 2005. A master regulator for biofilm formation by Bacillus subtilis. Mol Microbiol 55:739–749.
16. Oliveira NM, Martinez-Garcia E, Xavier J, Durham WM, Kolter R, Kim W, Foster KR. 2015. Biofilm Formation As a Response to Ecological Competition. PLoS Biol 13:e1002191.
17. O’Toole GA, Kolter R. 1998. Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a genetic analysis. Mol Microbiol 28:449–461.
18. Pratt LA, Kolter R. 1998. Genetic analysis of Escherichia coli biofilm formation: roles of flagella, motility, chemotaxis and type I pili. Mol Microbiol 30:285–293.
19. Prigent-Combaret C, Vidal O, Dorel C, Lejeune P. 1999. Abiotic surface sensing and biofilm-dependent regulation of gene expression in Escherichia coli. J Bacteriol 181:5993– 6002.
20. Schluter J, Nadell CD, Bassler BL, Foster KR. 2015. Adhesion as a weapon in microbial competition. ISME J 9:139–149.
62 Chapter IV
Conclusions and Future Directions
Parts of the text presented in this chapter were published in:
Frenkel ES, Ribbeck K. 2015. Salivary mucins in host defense and disease prevention. J
Oral Microbiol 7:29759.
63 The diseases that result from dysregulated mucin production indicate that mucins play a key role in maintaining health. For example, defects in mucin production can lead to ulcerative colitis when mucins are under produced or cystic fibrosis when mucins are overproduced (1, 2).
More specifically, in vivo studies using Muc5b knockout mice have shown that reduced production of Muc5b can lead to increased mortality due to infection (3). These examples indicate that mucins may be important for disease prevention in the oral cavity, but, prior to the work presented in this thesis, little was known about how MUC5B affects microbes to protect the oral cavity.
To understand how MUC5B could aid in the prevention of oral diseases, I study MUC5B in the context of one of the most common diseases – dental caries (4). My work focuses on two bacteria commonly found in the oral cavity, Streptococcus mutans, which can cause dental caries when it attaches to the tooth surface, and S. sanguinis, which is associated with healthy oral conditions (5, 6). Results show that MUC5B salivary mucins modulate the physiology of these bacteria in ways that reduce surface colonization and promote bacterial coexistence. The findings presented in this thesis illustrate that MUC5B likely plays a key role in protecting the oral cavity from disease and lay the foundation for future research that delves deeper into the complex relationship between mucins and the microbes that inhabit the oral cavity.
In Chapter 2, I show that MUC5B reduces S. mutans attachment and biofilm formation on glass and hydroxyapatite even in the presence of sucrose. MUC5B does not alter S. mutans growth or have bactericidal effects, therefore the reduction in surface colonization results from a shift in biofilm cells to the planktonic state. S. mutans is a biofilm-forming oral bacterium that creates dental caries when it attaches to and grows on tooth surfaces. As S. mutans metabolizes dietary sucrose, it produces acid as a by-product, which
64 decreases local pH leading to destruction of tooth surface. To better understand how MUC5B influences S. mutans, I determine if the reduction in surface colonization is mucin-specific or generalizable to other polymers that increase viscosity. To answer this question, I repeated surface colonization experiments with methylcellulose instead of MUC5B. Methylcellulose has been used in previous studies to mimic the viscosity of mucus, but does not have the same complex branched sugar chains as mucin (7, 8). I found that methylcellulose did not significantly reduce S. mutans attachment or biofilm formation, therefore viscosity is likely not causing the observed reduction in attachment and biofilm formation. Further experiments could be performed, however, using other polymers such as dextran and polyethylene glycol. If another polymer produced the same effect, this could indicate specific features of MUC5B that may be playing a role in the reduction of S. mutans surface colonization. Another key question is whether the observed reduction in bacterial surface colonization in the presence of MUC5B can be generalized to other mucins and microbes. The answer is most likely yes, since similar results have been shown for Muc5ac porcine gastric mucin, which reduces Pseudomonas aeruginosa and Candida albicans biofilm formation (9, 10).
To better understand how MUC5B would affect other aspects of bacterial physiology in a more complex environment, I developed a dual-species competition model with S. mutans and S. sanguinis in Chapter 3. In this model, I study bacterial viability to assess how MUC5B influences interspecies competition. Results show that MUC5B significantly increases S. mutans and S. sanguinis coexistence. This model represents a first step towards understanding how MUC5B influences the oral microbiota, but more complex models are needed to better recapitulate the environment of the oral cavity. Follow up studies could use a similar model with other oral bacteria and fungi or plaque from dental patients, which has been harvested and used
65 in several studies to gain an understanding of the oral microbiota in different disease states (11,
12). In vivo studies using hamster models have also been used to study methods for cavity prevention and could be used in this context to better understand how MUC5B modulates the oral microbiota (13, 14). For example, hamsters could be fed a high sucrose diet with or without mucin supplements then the composition of their oral microbiota could be evaluated using 16S
RNA sequencing.
After determining that MUC5B increases bacterial coexistence, I evaluated several aspects of S. mutans and S. sanguinis growth in the presence of MUC5B to better understand how MUC5B could be influencing competition. Results indicate that MUC5B could reduce interspecies competition by altering the cells’ mode of growth. Specifically, in the dual- species culture, MUC5B reduces the number of cells in the mixed-species biofilm, thereby increasing the relative number of cells in the less competitive planktonic state (15, 16). Although this dual-species competition model does not fully capture the complex environment of the oral cavity, it provides an initial understanding of how MUC5B could protect the oral cavity by preventing opportunistic or pathogenic microbes from becoming dominant species.
The work presented in this thesis provides a foundation for other studies that further characterize how MUC5B is able to reduce bacterial surface colonization and interspecies competition. Besides acting as a physical barrier, there are two primary mechanisms through which salivary mucins can interact with microbes to provide protection: 1) salivary mucins can agglutinate microbes, which would facilitate bulk removal during swallowing, and 2) salivary mucins can cause the dispersal of microbes. Because there are few studies indicating that S. mutans directly binds MUC5B, the second mechanism likely plays a larger role. The increase in
S. mutans dispersal illustrated in this thesis may be a generalizable mechanism that mucins use to
66 reduce microbial virulence since other studies in the Ribbeck lab have also shown that Muc5ac porcine gastric mucin promotes P. aeruginosa and C. albicans dispersal (9, 10). Mucins’ ability to reduce microbial virulence by keeping cells in an individualized state without killing the organism is a unique mechanism of protecting the body; it allows opportunistic pathogens to survive in the microbiota but limits their pathogenic potential. In contrast, many antibacterial proteins in the oral cavity, such as lysozyme, protect the body by killing harmful organisms (17).
One main question that warrants further investigation is whether the observed microbial phenotypes in the presence of MUC5B are caused by a physical effect or a change in bacterial gene regulation. An example of a physical effect that is not related to increased viscosity would be that the mucin polymer networks surrounding bacterial cells reduce diffusion of secreted factors, such as quorum sensing molecules, which are necessary for biofilm formation. For example, competence-stimulating peptides are secreted molecules that are known to play a role in S. mutans biofilm formation (18). These peptides have a prominent hydrophobic patch, which could facilitate mucin-peptide interactions (19). Another way MUC5B could influence bacteria is through specific glycan interactions, which could lead to downstream changes in genetic regulation that reduce microbial virulence. The ability of mucins to regulate gene expression has been demonstrated by several studies showing that intestinal and gastric mucins alter bacterial genetic regulation and by Dr. Kavanaugh in the Ribbeck who demonstrated that Muc5ac down- regulates C. albicans virulence traits (10, 20, 21).
There are several experimental strategies that could be used to further dissect the mechanism by which MUC5B causes S. mutans and S. sanguinis dispersal. First, a global gene analysis study, such as RNA-sequencing, could be performed to provide insight into changes in gene regulation that may take place when oral bacteria are exposed to salivary mucins. S. mutans
67 biofilms could be grown in medium with and without MUC5B, then cells from the supernatant and biofilm could be harvested separately. Comparing RNA expression in these four cell populations would indicate whether cells exposed to MUC5B have up- or down-regulated specific genes or pathways that lead to the observed reduction in surface colonization. If the genetic profiles of MUC5B and non-MUC5B exposed cells have similar genetic profiles, this would indicate that MUC5B likely alters bacterial surface colonization through a physical effect.
Another way to understand how MUC5B could be affecting S. mutans physiology is by studying how secreted molecules may change in the presence of MUC5B. Two important secreted molecules produced by S. mutans are competence-stimulating peptide, which plays a role in biofilm formation as mentioned above, and mutacin, which is important for interspecies competition (18, 22, 23). These molecules could be purified from S. mutans cultures grown in the presence and absence of MUC5B; a change in the amount of these molecules in the presence of MUC5B could indicate that MUC5B is influencing their production or directly interacting with them, which would decrease their effective concentration.
To better understand how oligosaccharides on MUC5B could be affecting S. mutans, specific saccharides could be grafted onto a polymer backbone then added to surface colonization experiments instead of MUC5B. Grafted oligosaccharides could be composed of a single type of saccharide or several different types in a configuration that mimics portions of oligosaccharide chains naturally found on MUC5B. For example, fucose has been identified as a common terminal sugar residue, and Lewis antigens are common terminal epitopes; both of these examples would be interesting to study and could provide insight into the effect of MUC5B sugars on oral bacteria (24, 25). If specific sugars or combinations of sugars reduce S. mutans surface colonization and interspecies competition, this would indicate that oral bacteria may
68 directly or indirectly interact with MUC5B sugar residues.
Because the oral cavity is easily accessible, determining if MUC5B levels change during disease states would be interesting and could provide insight into how MUC5B could be used as a diagnostic marker of disease. The oral cavity can be a mirror for changes that are happening in other parts of the body, therefore MUC5B could be an indicator of diseases that are not specific to the oral cavity. For example, one study shows that once an individual is infected with HIV-1, the concentration of MUC5B in whole saliva is significantly decreased compared with non- infected individuals, which could make MUC5B a diagnostic marker of HIV-1 infection (26).
Further research is needed to better understand how MUC5B is able to create a gel that houses millions of oral bacteria while coaxing potentially harmful microbes into passive existence. Understanding how mucins protect the body could open the doors to an entirely new set of therapeutic tools that aim to prevent microbes from transitioning into a pathogenic state as opposed to antibiotics, which treat the microbe once it is already virulent and can lead to antibiotic resistance. Based on the findings presented in this thesis, MUC5B could potentially be used as a useful therapeutic or preventative treatment for oral diseases like periodontal disease and Candidiasis that are caused by an overgrowth of opportunistic pathogens (27–29). In these cases, MUC5B could ideally be used to shift the oral microbiota back to a state of health.
Although the use of mucins as therapeutics is promising, manufacturing full-length mucin mimetics in high yields is a challenge that has only begun to be investigated. One potentially more tangible goal is to identify specific sugars or sugar motifs on mucins that induce the same phenotypic effects described in this research. Once the influence of mucins on microbes is fully characterized, salivary mucins or engineered mimetics could potentially be used as therapeutic tools to prevent or treat diseases in novel ways.
69 References
1. Rose MC, Voynow JA. 2006. Respiratory tract mucin genes and mucin glycoproteins in health and disease. Physiol Rev 86:245–278.
2. Van der Sluis M, De Koning B, De Bruijn A, Velcich A, Meijerink J, Van Goudoever J, Büller H, Dekker J, Van Seuningen I, Renes I, Einerhand A. 2006. Muc2-deficient mice spontaneously develop colitis, indicating that MUC2 is critical for colonic protection. Gastroenterology 131:117–29.
3. Roy MG, Livraghi-Butrico A, Fletcher AA, McElwee MM, Evans SE, Boerner RM, Alexander SN, Bellinghausen LK, Song AS, Petrova YM, Tuvim MJ, Adachi R, Romo I, Bordt AS, Bowden MG, Sisson JH, Woodruff PG, Thornton DJ, Rousseau K, De la Garza MM, Moghaddam SJ, Karmouty-Quintana H, Blackburn MR, Drouin SM, Davis CW, Terrell KA, Grubb BR, O’Neal WK, Flores SC, Cota-Gomez A, Lozupone CA, Donnelly JM, Watson AM, Hennessy CE, Keith RC, Yang IV, Barthel L, Henson PM, Janssen WJ, Schwartz DA, Boucher RC, Dickey BF, Evans CM. 2014. Muc5b is required for airway defence. Nature 505:412–416.
4. World Health Organization. Fact Sheet N318 - Oral Health. 2012.
5. Becker MR, Paster BJ, Leys EJ, Moeschberger ML, Kenyon SG, Galvin JL, Boches SK, Dewhirst FE, Griffen AL. 2002. Molecular analysis of bacterial species associated with childhood caries. J Clin Microbiol 40:1001–1009.
6. Caufield PW, Dasanayake AP, Li Y, Pan Y, Hsu J, Hardin JM. 2000. Natural history of Streptococcus sanguinis in the oral cavity of infants: evidence for a discrete window of infectivity. Infect Immun 68:4018–4023.
7. Ivic A, Onyeaka H, Girling A, Brewis IA, Ola B, Hammadieh N, Papaioannou S, Barratt CLR. 2002. Critical evaluation of methylcellulose as an alternative medium in sperm migration tests. Hum Reprod 17:143–149.
8. Smith DJ, Gaffney EA, Gadêlha H, Kapur N, Kirkman-Brown JC. 2009. Bend propagation in the flagella of migrating human sperm, and its modulation by viscosity. Cell Motil Cytoskeleton 66:220–236.
9. Caldara M, Friedlander RS, Kavanaugh NL, Aizenberg J, Foster KR, Ribbeck K. 2012. Mucin Biopolymers Prevent Bacterial Aggregation by Retaining Cells in the Free- Swimming State. Curr Biol 22:2325–2330.
10. Kavanaugh NL, Zhang AQ, Nobile CJ, Johnson AD, Ribbeck K. 2014. Mucins Suppress Virulence Traits of Candida albicans. mBio 5:e01911-14.
70 11. Yost S, Duran-Pinedo AE, Teles R, Krishnan K, Frias-Lopez J. 2015. Functional signatures of oral dysbiosis during periodontitis progression revealed by microbial metatranscriptome analysis. Genome Med 7:27.
12. Duran-Pinedo AE, Yost S, Frias-Lopez J. 2015. Small RNA Transcriptome of the Oral Microbiome during Periodontitis Progression. Appl Environ Microbiol 81:6688-99.
13. Smith DJ, Taubman MA, Ebersole JL. 1979. Effect of oral administration of glucosyltransferase antigens on experimental dental caries. Infect Immun 26:82–89.
14. Smith DJ, Taubman MA, Ebersole JL. 1982. Effects of local immunization with glucosyltransferase on colonization of hamsters by Streptococcus mutans. Infect Immun 37:656–661.
15. Oliveira NM, Martinez-Garcia E, Xavier J, Durham WM, Kolter R, Kim W, Foster KR. 2015. Biofilm Formation As a Response to Ecological Competition. PLoS Biol 13:e1002191.
16. Schluter J, Nadell CD, Bassler BL, Foster KR. 2015. Adhesion as a weapon in microbial competition. ISME J 9:139–149.
17. Edgerton M, Koshlukova SE. 2000. Salivary Histatin 5 and its Similarities to the Other Antimicrobial Proteins in Human Saliva. Adv Dent Res 14:16–21.
18. Li Y-H, Tang N, Aspiras MB, Lau PCY, Lee JH, Ellen RP, Cvitkovitch DG. 2002. A Quorum-Sensing Signaling System Essential for Genetic Competence in Streptococcus mutans Is Involved in Biofilm Formation. J Bacteriol 184:2699–2708.
19. Syvitski RT, Tian X-L, Sampara K, Salman A, Lee SF, Jakeman DL, Li Y-H. 2007. Structure-Activity Analysis of Quorum-Sensing Signaling Peptides from Streptococcus mutans. J Bacteriol 189:1441–1450.
20. Tu QV, McGuckin MA, Mendz GL. 2008. Campylobacter jejuni response to human mucin MUC2: modulation of colonization and pathogenicity determinants. J Med Microbiol 57:795–802.
21. Dhanani AS, Bagchi T. 2013. The expression of adhesin EF-Tu in response to mucin and its role in Lactobacillus adhesion and competitive inhibition of enteropathogens to mucin. J Appl Microbiol 115:546–554.
22. Kreth J, Merritt J, Shi W, Qi F. 2005. Competition and Coexistence between Streptococcus mutans and Streptococcus sanguinis in the Dental Biofilm. J Bacteriol 187:7193–7203.
23. Kreth J, Zhang Y, Herzberg MC. 2008. Streptococcal Antagonism in Oral Biofilms: Streptococcus sanguinis and Streptococcus gordonii Interference with Streptococcus mutans. J Bacteriol 190:4632–4640.
71 24. Klein A, Carnoy C, Wieruszeski JM, Strecker G, Strang AM, Van Halbeek H, Roussel P, Lamblin G. 1992. The broad diversity of neutral and sialylated oligosaccharides derived from human salivary mucins. Biochemistry (Mosc) 31:6152–6165.
25. Thomsson KA, Schulz BL, Packer NH, Karlsson NG. 2005. MUC5B glycosylation in human saliva reflects blood group and secretor status. Glycobiology 15:791–804.
26. Zhang N, Zhang Z, Feng S, Wang Q, Malamud D, Deng H. 2013. Quantitative analysis of differentially expressed saliva proteins in human immunodeficiency virus type 1 (HIV-1) infected individuals. Anal Chim Acta 774:61–66.
27. Kumar PS, Griffen AL, Barton JA, Paster BJ, Moeschberger ML, Leys EJ. 2003. New Bacterial Species Associated with Chronic Periodontitis. J Dent Res 82:338–344.
28. Paster BJ, Boches SK, Galvin JL, Ericson RE, Lau CN, Levanos VA, Sahasrabudhe A, Dewhirst FE. 2001. Bacterial diversity in human subgingival plaque. J Bacteriol 183:3770– 3783.
29. Kim J, Sudbery P. 2011. Candida albicans, a major human fungal pathogen. J Microbiol 49:171.
72